Abstract
After completing fertilization with fusion of the pronuclei during syngamy, the zygote now has a diploid complement of chromosomes, undergoes its first mitotic division and then continues to divide by mitosis into a number of smaller cells known as blastomeres. In humans, the first few cleavage divisions take place in the oviduct, before the embryo reaches its site of implantation in the uterus (Figure 5.1).
Preimplantation Development
After completing fertilization with fusion of the pronuclei during syngamy, the zygote now has a diploid complement of chromosomes, undergoes its first mitotic division and then continues to divide by mitosis into a number of smaller cells known as blastomeres. In humans, the first few cleavage divisions take place in the oviduct, before the embryo reaches its site of implantation in the uterus (Figure 5.1).
Figure 5.1 Development of the mammalian embryo. The oocytes released from the ovary (OV) enter the ampulla where they are fertilized (F) and then are transported along the fallopian tube, cleaving to generate the morula stage (M). The blastocyst(B) expands, hatches and then implants (I) in the endometrium (E) of the uterus (U).
In contrast to oogenesis, where the cell undergoes a period of growth without replication or division, early embryo cleavage involves intense DNA replication and cell division in the absence of growth; the overall size of the embryo does not change as a result of the early cleavage divisions. As cleavage progresses, the embryo polarizes, and differences arise between the blastomeres; this process of differentiation may be regulated by unequal distributions of cytoplasmic components previously laid down in the oocyte during oogenesis, or by changes occurring in the blastomeres as a result of new embryonic gene transcription during development (Figure 5.2). Each blastomere nucleus will be subjected to a different cytoplasmic environment, which, in turn, may differentially influence activity of the genome. As a result, after the onset of zygote gene activation and subsequent differentiation, eventually the blastomeres set off on their own specific program of development. Maternal mRNA encoding developmental information is essential in early differentiation and has been found to persist during specific patterns of gene expression until gastrulation in some species; embryos in some bats and marsupials can remain in a state of diapause (dormancy) within the uterine cavity for many months. In mammals, maternal mRNA rapidly disappears after the major activation of the genome, i.e., at the two-cell stage in the mouse, four- to six-cell in humans, and eight-cell in sheep and cattle. Localized short- and long-lived maternal mRNAs probably regulate the initial stages of differentiation; there is some evidence to suggest that stores of maternal RNA in the oocytes of older women may be depleted, perhaps due to dysfunction or disruption of the mechanisms that control its storage.
Figure 5.2 Timescale of early human embryo development up to the blastocyst stage, correlating morphological changes with developmental events. Time is in hours.
Genome Activation
As described in Chapter 3, the developing oocyte accumulates reserves of mRNA, proteins, organelles, etc. These maternal transcripts and proteins are required to support and direct early development, and are progressively degraded as the new embryonic genome is increasingly transcribed. In vitro, the early embryo shows very little metabolic activity during its first few cleavage divisions. Stored maternal mRNA directs the first two cleavage divisions, and then activation of the new embryonic genome provides novel transcripts and reprograms the pattern of gene expression to direct further development. After the long period of gene suppression in both gametes, the new embryonic cell cycle must be precisely timed and regulated, with a correct timing of DNA synthesis during S-phase. The cycle during which the zygote genome is activated is always the longest cell cycle of preimplantation development: any delay at this time may cause the level of mRNA to fall below a critical threshold, and without appropriate zygotic genome activation (ZGA), the mammalian embryo fails to develop further. This critical transition takes place during the four-to eight-cell stage in humans, and maternal mRNA rapidly disappears whilst the zygote genome gradually increases its expression. However, the transition is not absolute; maternal transcripts are degraded at different rates, and the stability of the maternally encoded proteins vary, so that some maternal information may still contribute to embryonic development after the embryonic genome is activated. A small amount of maternal message is needed almost up until the blastocyst stage; therefore, previous failures at any stage of oocyte development, maturation and handling can affect development even after ZGA. Gene expression involves conformational changes in nucleosome organization (like uncoiling a spring), regulated by interactions among DNA methylation, histone acetylation and messenger RNA polyadenylation patterns. Activation of the embryonic genome follows a series of progressive steps; the timing and coordination of gene expression can be regulated at the level of maternal mRNA translation. The newly formed embryonic genome undergoes extensive epigenetic modifications that result in a program of gene expression that is highly regulated. Any disruption during these initial steps can have a far-reaching impact on embryo development.
1. Maternal RNA transcripts are depleted, and new embryonic mRNA is transcribed. The replacement of maternal transcripts by those of the zygote occurs at different rates for different genes; this may be due to the relative importance of different transcripts for immediate developmental events, or because the protein products differ in their stability, or both. The expression of approximately 1800 mRNAs is modulated during the first 3 days of development; the majority are downregulated or destroyed, a small number are upregulated on days 1 and 2, and a large group of mRNAs become increasingly abundant on Day 3 (Dobson et al., 2004); these may represent preferentially stable mRNAs in the pool of maternal mRNA that is being degraded.
2. There is a qualitative shift in protein synthesis and in post-translational modification.
3. A functional nucleosomal structure develops, the nuclear organizing region (NOR).
Establishing the precise timing of genome activation is related to the sensitivity of methods available for its detection; new protein synthesis activity has been detected at different stages in different species:
Recent transcriptome analysis has shown that gene expression during ZGA is carefully orchestrated: genes involved in transcription and RNA metabolism are highly expressed (Hamatani et al., 2004; Wang et al., 2004; Zeng & Schultz, 2004), and most of the newly synthesized ZGA transcripts are quickly translated (Flach et al., 1982). Epigenetic chromatin remodeling has been proposed as the main mechanism regulating the ZGA: epigenetic marks involve post-translational modifications of nucleosomal histones (methylation, acetylation, phosphorylation and ubiquitination), DNA methylation and non-histone proteins that bind to chromatin.
There is evidence to suggest that a minor degree of transcriptional activity does take place prior to the major activation of the genome, with a period of minor gene activation from the paternal pronucleus in the one-cell embryo, followed by a period of gene activation in the two-cell embryo when maternal mRNA and zygotic gene transcripts are handled differently, so that transcription and translation of nascent transcripts is delayed (Wiekowski et al., 1991; Nothias et al., 1995; Schultz, 2002). In human preimplantation embryos, reverse transcriptase-polymerase chain reaction (RT-PCR) detected early transcripts for two paternal Y chromosome genes, ZFY and SRY. ZFY transcripts were detected at the pronucleate stage, 20–24 hours after in-vitro insemination, and at intermediate stages up to the blastocyst stage. SRY transcripts were also detected at two-cell to blastocyst stages (Ao et al., 1994).
In 1938, the Carnegie Institution of Washington funded a 7-year study with the aim of finding and characterizing early embryos recovered from fertile married women undergoing therapeutic hysterectomy. Over a period of 15 years, Arthur Hertig and John Rock examined excised fallopian tubes and uteri, allowing them to describe 34 embryos aged up to 17 days post-fertilization: the earliest were at the two-cell stage, and the latest were undergoing gastrulation (Hertig et al., 1954, 1956). Their analyses led them to make several key observations about the dynamics of fertilization and early embryo development, as well as providing insight into rates of embryo loss after natural conception.
Cleavage increases the number of nuclei, which amplifies the number of templates that will facilitate the production of specialized proteins needed for the later processes of compaction and differentiation to the blastocyst stage. As mentioned above, changes in chromatin structure, rather than changes in the activity of the transcriptional apparatus, may underlie the timing and basis for ZGA. Transcription factors must be available that can bind to the DNA, and a functional physical structure, the NOR develops, producing conformational changes in the DNA structure that will allow the binding of promoters and enhancers of transcription. In the mouse and rabbit, there is a general chromatin-mediated repression of promoter activity. Repression factors are inherited by the maternal pronucleus from the oocyte, but are absent in the paternal pronucleus; they become available sometime during the transition from a late one-cell to a two-cell embryo (Henery et al., 1995). This means that paternally inherited genes are exposed to a different environment in fertilized eggs than are maternally inherited genes, a situation that could contribute to genomic imprinting.
The formation of the NOR is related to the nuclear:cytoplasmic ratio. A titratable factor in the cytoplasm, possibly related to cdc25, may be diluted with the increase in nuclear:cytoplasmic ratio, driving maturation promoting factor (MPF) and a kinase cascade that triggers mitosis. The gradual depletion of cdc25 causes a pause in cleavage, which allows time for the NOR to develop, and mitosis induces a general repression of promoters prior to initiation of zygotic gene expression. Enhancers then specifically release this repression. A biological clock may delay transcription until both paternal and maternal genomes are replicated; they must then be remodeled from a postmeiotic state to one in which transcription is repressed by chromatin structure. The chromatin structure must have a configuration that allows specific transcription enhancers to relieve/reverse repression at appropriate times during development. Differential hyperacetylation of histone H4 (particularly on DNA in the male pronucleus) has been implicated in the remodeling of maternal and paternal chromatin, and depletion of maternally derived histones has also been suggested as one of the mechanisms involved in ZGA (Adenot et al., 1997).
Chromatin-mediated repression of promoter activity prior to ZGA is similar to that observed during Xenopus embryogenesis; this mechanism ensures that genes are not expressed until the appropriate time in development. When the time is right, positive factors such as enhancers can begin their activity. The mechanism by which enhancers communicate with promoters seems to change during development and may depend upon the presence of specific co-activators. In the mouse, ZGA occurs during the second cell division cycle (two-cell stage) and seems to be regulated by a ‘zygotic clock’ that measures the time following fertilization rather than progression through the first cell cycle. There is evidence that circadian clock genes may be involved in programming an appropriate timing, so that development is synchronized with endocrine and other local factors to ensure successful implantation. In support of this concept, expression of circadian clock genes has been found in the reproductive tract and conceptus of the mouse during the first 4 days of pregnancy (Johnson et al., 2002). It may be that in vivo, mammalian ZGA is a time-dependent mechanism that must interact in synchrony with the cell cycle and other physiological events.
In human embryos, the major wave of genome activation occurs on Day 3, independent of cell number: embryos that arrest with fewer than eight cells still show evidence of ZGA (Dobson et al., 2004). This corresponds to the first wave of ZGA in mice, at 26–29 hours post fertilization (Vassena et al., 2011). Further changes in morphology, i.e., compaction and cavitation, also depend on timing of development, rather than cell number – irrespective of lysis or fragmentation of one or more blastomeres. Inherited maternal/paternal factors have a significant influence on zygote development, including those that mediate RNA metabolism/translation and cytokinesis as well as ploidy and epigenetic factors. During further development, the embryo’s morphology, physiology and metabolism are shaped by an interplay between its genotype and its response to environmental factors. The human embryo is exquisitely sensitive to environmental signals and shows a high degree of plasticity by modulating its metabolism, gene expression and rate of cell division. This developmental plasticity, with effects on gene methylation and expression, has the potential to influence later health and well-being in postnatal life (Bateson et al., 2004; Rosenbloom, 2018).
Imprinting
During oogenesis and spermatogenesis, the maternal and paternal chromosomes are packaged in a manner that affects subsequent transcription of some of the genes during development. The DNA sequence that specifies the parental genes is not altered, but the way in which it is chemically modified and packaged in chromatin affects the expression of the genes. Because the genetic code itself remains the same, this modification is referred to as an epigenetic change, and the phenomenon is known as imprinting. The pattern of epigenetic change is parentally specific, i.e., the genes affected (imprinted) in oocytes differ from those imprinted in sperm – there is differential expression of the two parental alleles of a gene. In terms of function, this means that although the oocyte and the sperm each contribute a complete set of genes, each set on its own is not competent to direct a complete program of development; a fully functional genome requires the combination of both paternal and maternal genes. The process of genomic imprinting is established during gametogenesis, and the nucleus of the zygote has an imprint memory that is retained by the embryo into both prenatal and postnatal life. Imprinting is highly regulated during preimplantation development; the topic is thus highly significant and relevant to in-vitro manipulations and culture, and will be discussed in detail in Chapter 15.
Compaction
The first event that determines the directed development of previously undifferentiated blastomeres is compaction. During the first few cleavage divisions to reach the four- to eight-cell stage, individual blastomeres can be clearly seen in the developing embryo. At about the third cleavage division there is a significant increase in RNA and protein synthesis, a marked change in the patterns of phospholipid synthesis, and the embryo undergoes compaction to form a morula. During this transition as the embryo is compacting, it must also make fundamental decisions regarding cell position, polarity and fate. The process is calcium-dependent, and requires prior transcription of the zygote genome. With compaction, the blastomeres flatten against each other and begin to form junctions between them, so that the boundaries between blastomeres can no longer be distinguished. The cells of the compacted embryo become highly polarized and tightly associated, with redistribution of surface microvilli and other plasma membrane components.
Coordination of these complex developmental processes requires communication between the cells; two types of intercellular junction have been described:
1. Structural tight junctions and desmosomes anchor the cells together and form an impermeable epithelial barrier between cells. Tight junctions are composed of several integral and peripheral proteins, including occludin and cingulin (ZO-1).
2. Low-resistance junctions such as gap junctions allow the flow of electrical current and the direct transfer of small molecules, including metabolites and second messengers (cAMP) between blastomeres.
Compaction has been extensively studied in the mouse: the distribution of dense microvillar and amicrovillar regions indicates surface polarity, and the distribution of endocytotic vesicles and actin filaments, as well as the location of the cell nucleus, demonstrates polarity in the cytoplasm. In the mouse, isolated blastomeres that are decompacted experimentally maintain their polarity, and compaction does not require either a prior round of DNA replication or protein synthesis (Kidder and McLachlin, 1985). Therefore, the four-cell embryo probably contains some of the proteins required for compaction. Although the factors that trigger the timing of its onset are not known, experimental evidence suggests that this may be regulated by post-translational modification of specific proteins such as E-cadherin. E-cadherin protein (uvomorulin) is expressed in the oocyte, and during all stages of preimplantation development. It is uniformly distributed on the surface of blastomeres and accumulates in the regions of intercellular contact during compaction. E-cadherin phosphorylation can be observed in the mouse eight-cell embryo. Culturing embryos in calcium-free medium inhibits E-cadherin phosphorylation and prevents compaction, but the situation is complex, and precise mechanisms behind the molecular basis for compaction and its timing remain unclear.
In human embryos, tight junctions begin to appear on Day 3, at the 6- to 10-cell stage, heralding the onset of compaction. The surface morphology of human oocytes and embryos has been studied with scanning electron microscopy (Santella et al., 1992; Dale et al., 1995; Nikas et al., 1996), and this showed that unfertilized oocytes 1 day after insemination were evenly and densely covered with long microvilli. The length and density of microvilli appeared to decrease in fertilized oocytes, and a further decrease was observed in Day 2 and Day 3 embryos with 2–12 cells. There was no evidence of surface polarity until Day 4, when it was evident in the majority of embryos with 10 or more cells. The microvilli appeared dense again with a polarized distribution over the free surface of the compacted blastomeres.
In the mouse, gap junctions are expressed at the eight-cell stage, and their de novo assembly during compaction is a time-dependent event. Inhibition of DNA synthesis during the third and fourth cell cycles has no effect on the establishment of gap junctional coupling during compaction (Valdimarsson and Kidder, 1995), but a delay of 10 hours in DNA synthesis during the second cell cycle results in the failure of gap junctional coupling at the time of compaction.
In human embryos, gap junctions are not apparently well developed until the early blastocyst stage, when intercellular communication is clearly seen between inner cell mass (ICM) cells (Figures 5.3 and 5.4; Dale et al., 1991).
Figure 5.3 Functional expression of gap junctions in the early human embryo shown by micro injection of the low molecular weight tracer lucifer yellow. There is no dye spread in four cells (a), ten cells (b) or morula (c). However, transfer occurs in the blastocyst stage (d). icm = inner cell mass.
Figure 5.4 Transmission electron micrographs showing tight junctions (TJR) and gap junctions (G) in the human morula. (a) is a section at the apical level between two polar cells at a magnification of ×75 500; (b) shows a typical section between a polar and an apolar cell at ×158 000. Arrowheads show sites of tight membrane contact.
Following compaction, the developing embryo is described as a morula, seen in the human normally 4 days after fertilization. The embryo now shows a significant increase and change of pattern in RNA, protein and phospholipid synthesis, and this results in a process of differentiation so that cells are now allocated to an ICM, with outer cells forming an epithelial layer of trophectoderm. Whereas early blastomeres are totipotent (as evidenced by experimental embryo splitting and chimera formation), at compaction the cells polarize radially, and differential division across this axis creates different populations of cells, with unequal distributions of organelles:
1. Outer polar cells with surface microvilli and redistribution of other plasma membrane components are restricted to the free outer surface of the embryo, and these cells form the trophectoderm.
2. Inner apolar cells with tight junctions containing basal nuclei, which will form the ICM.
These morphological transitions are thought to be brought about by differential gene expression with corresponding protein expression profiles, but these have not yet been clearly defined at the molecular level. Experimental interference with adhesion between cells during compaction shows that this process is important in determining cell lineages. ICM cells preferentially communicate with each other and not with trophectoderm cells via gap junctions, whereas trophectoderm cells communicate with each other and not with ICM cells. The ratio of trophectoderm to ICM cells can be influenced by culture conditions in vitro, and this might have implications for further embryonic/fetal development.
Cavitation
Between the 16- and 32-cell stage, a second morphological change occurs, known as cavitation. The trophectoderm is the first cell to differentiate after ZGA, and these polarized blastomeres compact to construct functional complexes of junctions and systems that can then transport ions and water to fill the blastocoelic cavity with fluid. Activation of Na+, K+ ATP-ase systems in the trophectoderm cells results in energy-dependent active transport of sodium pumped into the central area of the embryo, followed by osmotically driven passive movement of water to form a fluid-filled cavity, the blastocoele. The movement of other ions such as chloride and bicarbonate also contributes to blastocoele formation. Immunohistochemistry shows that the trophectoderm cells are sitting on a basement membrane, and tight junctions form a continuous belt between trophectoderm cells, preventing leakage of small ions in the blastocoelic fluid.
Blastocoele formation and expansion is critical for further development, as it is essential for further differentiation of the ICM. This is now bathed in a specific fluid medium, which may contain factors and proteins that will influence cell proliferation and differentiation. The position of cells within the ICM in relation to the fluid cavity might also contribute to the differentiation of the outer cells into primitive endodermal cells.
Apoptosis can be seen at the blastocyst stage, localized to the ICM: this may represent a mechanism for the elimination of inappropriate or defective cells.
The trophectoderm cells will eventually form the placenta and extraembryonic tissue. Myxoploidy of trophectoderm cells is a common feature in all animal species, regardless of their implantation mechanisms; mouse and cow, which differ completely in their mechanism of implantation, show this feature, with chromosome complements of 2n, 4n, 8n in their trophectoderm cells. However, in humans it could possibly be considered as the initiation of syncytiotrophoblast formation. The regulation of this process, and apparent lack of division in these cells, remains a mystery – but it seems to be related to the appearance of giant cells in the trophectoderm, suggesting that regulation of the nuclear/cytoplasmic ratio is involved. It is interesting to note that there is a counterpart of ‘giant cells’ in the uterus around the time of implantation. A retroviral syncytin envelope gene with cell–cell fusion activity has been identified in mammalian syncytiotrophoblast, and is postulated to be responsible for syncytiotrophoblast formation (Heidmann et al., 2009).
Blastocyst Expansion and Hatching
The embryo is enclosed in the zona pellucida during these early stages of development, keeping the cells together prior to compaction and acting as a protective barrier. If the ICM divides at this early stage, monozygotic (identical) twins may develop.
In humans, the early blastocyst (Day 4/5) initially shows no increase in size, and a cavity representing <30% of its volume is visible. It subsequently expands over the next 1 or 2 days (Day 5/6) by active accumulation of fluid in the central blastocoelic cavity, which expands to form 70% of the embryo volume. Blastocyst expansion is driven by the trophectoderm, as described above.
Time-lapse imaging reveals that expansion is a dynamic process, with pulses of oscillating contraction/blastocoel collapse/recovery at intervals of 2 to 4 hours (Huang et al., 2016). Variations between different embryos can be seen, particularly at later times during the periods of expansion. The data further suggest that periodicity and amplitude of oscillations may be modulated by the zona pellucida: embryo-specific variations in expansion kinetics may reflect variations in the zona. Rates of blastocyst expansion were found to correlate with viability/implantation, with stronger contractions related to impaired zona hatching, i.e., inadequate recovery from blastocoel contraction may jeopardize hatching and subsequent implantation. Blastocyst expansion could thus represent an in-vitro ‘stress test’ of embryo viability, confirming that this feature is a useful tool in selection of embryos for transfer (see Chapter 11).
Before the blastocyst can start the process of implantation, it must free itself from the protective zona pellucida, which becomes visibly thinner as the late blastocyst expands; in vitro, initiation of the hatching process can be seen as trophectoderm cells ‘escaping’ from the zona pellucida (Figure 5.5). Hatching is completed within a few hours, and the freed blastocyst is separated from its empty zona (see also Chapter 11, Figure 11.8).
Figure 5.5 Scanning electron micrograph of a hatching human embryo. The microvilli on the surface of the trophectoderm cells are bared owing to internal pressure in the blastocoel and dissolution of the zona pellucida.
Cell Fate and Cell Lineages
The fate of each blastomere is influenced by mechanisms that achieve a balance between pluripotency and differentiation; the expression of lineage-specific genes differs between different species. At the earliest stages of development, the transcriptional machinery that will direct differentiation is not switched on, and transcription is under the control of specific transcriptional regulators, regulatory RNAs and chromatin remodeling machinery, which are in turn influenced by epigenetic marks, cell positional history, cell polarity and orientation of division. As discussed in Chapter 3, the mammalian oocyte has evidence of polarity, and this has been confirmed in human oocytes. Maternal factors are subsequently important in establishing polarities, regulating cleavage planes and in allocating specific blastomeres to their eventual fate. By the morula stage, cell fate decisions have been made, and an axis is established with embryonic (ICM) and abembryonic (trophectoderm) poles. Cells positioned on the inside of the morula retain pluripotency, and those on the outside develop into extraembryonic trophectoderm which will support the development of the embryo in the uterus and influence embryonic patterning before gastrulation via signaling mechanisms. The generation of inside cells requires outer cells to divide in an orientation such that one daughter cell is directed inwards during the 8- to 16-cell and 16- to 32-cell stages; these divisions are known as differentiative, in contrast to conservative divisions in which both daughter cells remain on the outside. Because inside and outside cells will follow different fates, differentiative divisions probably distribute cell fate-determining factors asymmetrically between the daughters. Several molecules that influence polarization have been identified: the Ca2+-dependent E-cadherin molecule is implicated in generating blastomere polarity, localized geographically at division together with the actin microfilament stabilizing protein ezrin. Homologues of PAR (partitioning defective proteins) also influence the regulation of cell polarization and the control of asymmetric cell divisions via positioning effects on the spindle. The transcription factor Cdx2 is required for the commitment of outer cells to the trophoblast (see Chapter 7).
The molecular basis for the generation and stabilization of polarity in development is not fully understood, but evidence is accumulating to suggest that cell fate may be determined as early as the four-cell stage in human embryos: mRNAs specific for a trophectoderm lineage (beta-hCG) were identified in a single blastomere of a four-cell embryo, and not at the two-cell stage; a single putative trophectodermal precursor appears to emerge during the second cleavage division (Hansis et al., 2002; Edwards and Hansis, 2005). In mouse embryos, lineage tracing experiments using fluorescent tracers and optional sectioning indicate that differences in developmental properties of individual blastomeres may be determined at the two-cell stage (Piotrowska et al., 2001; Piotrowska-Nitsche et al., 2005). These experiments showed that the first cell to divide to the four-cell stage contributed preferentially to the embryonic cell lineage, whereas the later-dividing blastomere contributed to the abembryonic (trophectoderm) lineage. By the blastocyst stage, the position of cells within regions of the blastocyst has an influence on their subsequent fate in postimplantation development (Gardner, 2001, 2007).
Although transcription in human blastomeres has been found to be similar up to the precompaction eight-cell stage, it is possible that they are already programmed to express lineage-associated genes (Galan et al., 2010; Wong et al., 2010). Single-cell RNA sequencing and analysis of protein distribution in human and mouse embryos from the zygote to the eight-cell stage revealed transcriptional programs that are conserved between the species, as well as some that are specific to human embryos (Blakeley et al. 2015). In a further study, CRISPR-Cas9 genome editing that targeted the OCT 4 gene (POU5F1) in mouse and human zygotes revealed that the loss of OCT 4 affects human and mouse embryos differently: the targeted human embryos initiated blastocyst formation, but the ICM formed poorly, and the embryos subsequently collapsed. The mutation downregulated both extraembryonic trophectoderm genes (CDX2) and those that regulate the pluripotent epiblast, including NANOG. OCT4-targeted mouse embryos continued to blastocyst development and maintained their expression of genes such as NANOG (Fogarty et al., 2017). These studies suggest that OCT4 has a different function in humans than in mice and is required earlier in human blastocyst development: its expression can be detected during cleavage and morula stages.
The ICM is a transitory state, and before the blastocyst implants into the uterine wall, its cells diverge into either early epiblast (E) or hypoblast/primitive endoderm (PE, see Chapter 6). The mechanisms of segregation have been extensively studied in the mouse, and Roode et al. (2012) observed that human embryos segregate PE by mechanisms that differ from those identified in mouse, rat and cow embryos, suggesting that the specification of cell lineages is different in different species. Glinsky et al. (2018) identified multilineage precursor cells emerging in human embryos at the morula stage on Day 4 that were sustained up to Day 7 of development. These cells were described as having a Multilineage-Markers-Expression phenotype (MLME cells); they suggest that lineage specification thus begins prior to the ICM stage, and that MLME cells congregate in the ICM, where they continue differentiation into more specialized cell types.
It is important to recognize that each step in the creation of a blastocyst is dependent upon the previous step, with new avenues for interaction between the cell populations leading to ever-increasing complexity of the embryo. In vitro, manipulations and culture environment have the potential for disrupting the highly sophisticated and carefully orchestrated events necessary for normal development and implantation.
Preimplantation Development in Mouse and Human Embryos
Although mouse and human embryos appear similar in morphology during their early post-fertilization development, key molecular differences in gene expression patterns and in developmental timing have been identified (Niakan & Eggan, 2013). The centrosome structure is contributed by the oocyte in the mouse, and by the sperm cell in humans. The pattern of DNA methylation and X-inactivation also differs between the two species (Fulka et al. 2004). Human embryos are far more susceptible to genetic instability than are mouse embryos, and numerous studies have revealed significant differences in cell cycle regulation, control of apoptosis and cytokine expression. Some of the established differences are summarized in Table 5.1 and Figure 5.6.
Table 5.1 Comparison of human and mouse preimplantation development
| Parameter | Human | Mouse |
|---|---|---|
| Average cell cycle time | 13–16 hrs | 10 hrs |
| Average embryo diameter | 140 µm (Volume = 8× that of mouse) | 70 µm |
| Zygotic gene expression | 4–8 cell stage, D3 | 2-cell stage, D1–2 |
| Waves of transcription | MII – 4c, maternal genes only 8c-blastocyst: maternal genes downregulated, zygotic genes upregulated | 2-cell: genes required for transcription & RNA processing 4–8 cell: genes involved in morphology & function |
| Compaction | 8–16 cell stage, D4 | Late 8-cell stage, D3 |
| Cavitation | From 35 cells, D4 to early D5 | From 32 cells, D3.25 |
| Blastocyst formation | 108–136 hrs; 64–100 cells, late D5 128–256 cells, early to late D6 >256 cells | 84–96 hrs; <64 cells D3.5 >64 cells D3.75 to D4.25 ~164 cells |
| Oct4 expression | 7-8c, D3; required for blastocyst formation Persistently expressed in TE, downregulated in a subset of PE cells, restricted to ICM on D6 | 4c, D2; downregulated in TE by D4.5, not required for blastocyst formation |
| GATA2 & 3 expression | Trophoblast of late blastocysts | Cleavage stage in all cells, then restricted to outer trophoblast in late blastocysts |
| Segregation of epiblast/hypoblast from ICM | D7; not dependent on FGF signaling | D4.5; dependent on FGF signaling, involves NANOG and Gata6 |
| Epiblast morphology | Flattened bilaminar disc | Cup-like egg cylinder |
| Implantation | Between D6 and D8 TE cells form invasive cytotrophoblast (CT) cells; CT cells proliferate & differentiate in placental villi: Multinucleated syncytial cells Extravillous trophoblast cells that invade uterine decidua | Day 4–4.5 Minimal early invasion; TE cells form early proliferative cells adjacent to the epiblast Polyploid trophoblast giant cells are budded off through endoreduplication, giving rise to extraembryonic ectoderm |
| Stem cell doubling time | 30–35 hrs | 12–15 hrs |
| Maintenance of stem cell lines | Does not require LIF | LIF-dependent |
Figure 5.6 Comparison of human and mouse peri-implantation development. Left panel: early human postimplantation development, with syncytiotrophoblast and primary villi. Right: early mouse postimplantation development, with extraembryonic ectoderm and ectoplacental cone.
Causes of Early Embryo Arrest
Cleaved embryos do frequently arrest their development in culture, and a great deal of research has been carried out in animal systems to elucidate possible causes and mechanisms. Studies conducted over the past 15 years continue to reveal that cultured embryos show significant changes in their patterns of gene expression. These changes affect not only survival during the preimplantation period but may also affect the ability of the embryo to implant and continue through normal fetal development. They may also have an impact on post-partum health and susceptibility to disease in later life. Understanding the mechanisms and signaling pathways involved in the embryo’s response to culture environments is of paramount importance in minimizing potential hazards to normal human preimplantation development in vitro. Differences observed in gene expression patterns between cultured and in-vivo-derived embryos may be due to changes in gene transcription, or to changes in mechanisms that cells use to regulate mRNA stability and half-life.
The longest cell division cycle during development is that during which genome activation takes place, when maternal transcripts are degraded and massive synthesis of embryonic transcripts is initiated. Maternal reserves are normally sufficient until transcription begins, but epigenetic effects of defective sperm can lead to accumulation of delays, with resulting arrested development. Antisperm antibodies can have deleterious effects at this stage, by immunoneutralization of proteins that signal division (CS-1) or regulation (Oct-3). After genome activation, the next critical stage is morula/blastocyst transition. Complex remodeling takes place, and poor sperm quality can compromise this transition (see Ménézo and Janny, 1997).
Embryonic arrest is frequently a result of events surrounding maturation but can be a result of any metabolic problem. In bovine and pig oocytes, insufficient glutathione inhibits decondensation of the sperm head and polar body formation, and genetic factors regulate the speed of preimplantation development. Genetic factors implicate enzyme deficiencies or dysfunctional regulation, which may have deleterious effects. In domestic animals, as in humans, there is an age-related maternal effect. Maternal age has an effect on embryo quality, especially on blastocyst formation – this may be related to an ATPase-dependent Na+/K+ pump mechanism, or to a poor stock of mRNA, poor transcriptional and/or post-transcriptional regulation or accelerated turnover of mRNA.
In clinical IVF, detection of two pronuclei is regarded as evidence that normal fertilization has taken place, and the formation of a normal mitotic spindle following fertilization is critical in order to ensure correct chromosomal alignment. Mistakes at this stage can be lethal, resulting in chromosomal disorders such as aneuploidy. In some cases a first cleavage division takes place when no pronuclei have been detected during the previous 24–28 hours, a phenomenon that has been described as ‘silent fertilization.’ The first cell division tends to be asymmetrical, and the embryos arrest during cleavage. Van Blerkom et al. (2004) undertook a multi-year study of oocytes and embryos in which silent fertilization was suspected, using scanning laser confocal fluorescence microscopy to study chromosomal and microtubular structures. They were able to visualize maternally and paternally derived spindles in embryos that had shown no sign of pronuclear evolution after multiple, closely spaced inspections at the 1-cell stage, with maternal and paternal spindles well separated. The authors suggest that the evolution of such embryos may have an unusual pattern of chromosomal segregation, leading to micro- or multinucleation. The mechanisms involved in silent fertilization could be due to defects in normal calcium signaling, inadequate cytoplasmic maturity or delayed release of sperm-derived factors that also modulate calcium signaling.
Fluorescent in-situ hybridization (FISH) analysis of cleaved human embryos has confirmed that chromosomal aberrations are found in a significant proportion of embryos which develop with regular cleavage and morphology; this undoubtedly contributes to the high wastage of embryos in human IVF.
Paternal Factors
Sperm quality may have an influence on embryogenesis and implantation potential. Increasing paternal age is thought to have an influence on fertility, possibly through increased nondisjunction in the sperm. Damage during spermatogenesis may be induced by reactive oxygen species and defective oxidative phosphorylation, or via inherited dysfunctional mitochondrial DNA. Fertilization by a sperm that is diploid, with incomplete decondensation and DNA activation or inadequate chromatin packaging, may cause aneuploidy or lack of genome competence in the embryo. The quality of condensation and packaging of sperm DNA are important factors for the initiation of human embryo development, even after ICSI. The centrosome, involved in microtubular organization, is the first epigenetic contribution of the sperm, and correct and harmonious microtubule arrangement is necessary for chromosome segregation and pronuclear migration. An abnormal sperm carrying an imperfect centrosome can disrupt mitosis, provoking problems at the beginning of embryogenesis with the formation of fragments, abnormal chromosome distribution and early cleavage arrest. Up to 25% of apparently unfertilized eggs may show signs of having initiated fertilization, but then have anomalies that prevent cell division. In bulls, there is a positive correlation between sperm aster formation at the time of fertilization and the bull’s fertility. In the human, paternal Y-linked genes are transcribed as early as the zygote stage, and compromised paternal genetic material could be transcribed at even this early stage, causing fertilization failure or embryonic arrest. Finally, as discussed in Chapter 3, spermatozoa lacking in or with defective oocyte activating factor may only partially activate oocytes and lead to abortive development.
Metabolic Requirements of the Early Mammalian Embryo in Vitro
Maternal oocyte reserves that are stored during the maturation process must be capable of supporting metabolism in the early embryo prior to ZGA. Genes coding for all of the required enzymes must be expressed with correct timing and in the appropriate equilibrium. There must also be an effective mechanism for repairing damaged DNA in the oocyte and sperm; an estimated 1.5 to 2 million repair operations are performed during the first cell cycle, and these are dependent upon adequate maternal reserves of mRNA, proteins, organelles, etc.
When the oocyte is fertilized and starts the process of transcription, the new embryo must maintain equilibrium between many different parameters:
1. The endogenous pool of metabolites, largely the result of final stages of oocyte maturation
2. Metabolic turnover of RNA messengers and proteins
3. Active uptake of sugars, amino acids and nucleic acid precursors
4. Passive transport, especially of lipids
5. Incorporation of proteins such as albumin, which can bind lipids, peptides and catecholamines.
Although maternal reserves play an important role, the environment of the embryo is also critical. Culture media rarely reflect in-vivo conditions: the efficacy of transport systems into the oocyte and early embryo must be taken into account. Culture conditions have a direct impact on transcription and translation; embryonic metabolism may be depressed, protein turnover accelerated and mitochondrial function may be impaired. Suboptimal culture conditions have been shown to decrease cell numbers and jeopardize embryo viability.
ATP as an energy source is a basic requirement, and mammalian cells can generate ATP either by aerobic oxidation of substrates to carbon dioxide and water, or by anaerobic glycolysis of glucose to lactic acid. Under in-vitro conditions, oocytes and embryos generate ATP by aerobic oxidative metabolism of pyruvate, lactate, amino acids and possibly lipids. These metabolites have been shown to be important prior to genomic activation; pyruvate can also remove toxic ammonium ions via transamination to alanine.
Pentose Phosphate Pathway, NADPH and Glutathione
An important feature of early embryo metabolism is linked to the activation process that is induced by sperm entry, which increases glycolysis and glucose uptake through transporters. These may provide energy by generating ATP, but upregulation of the pentose phosphate pathway (PPP) at the time of pronuclear formation is a more significant metabolic parameter during this period. The PPP generates ribose 5-phosphate, a nucleotide precursor for subsequent DNA synthesis and replication. Upregulation of glucose metabolism via the PPP requires a fully grown pronucleus; the activity of the PPP influences the onset of the first S-phase in both male and female pronuclei, and continues to influence embryo development up to the blastocyst stage. The PPP also generates NADPH, which is involved in the majority of anabolic pathways: 1 mole of glucose 6-phosphate generates 1 mole of ribose 5-phosphate plus 2 moles of NADPH. NADPH further allows methionine to be recycled from homocysteine, with the formation of folic acid via methylene tetrahydrofolate reductase (MTHFR) (see Chapter 1: The 1-Carbon Cycle). This pathway influences imprinting processes in the oocyte, and is also involved in thymidine synthesis (5-methyl-uracyl, see section Vitamins). NADPH is also required to reduce oxidized glutathione (GSSG). The synthesis of glutathione from cysteine is energy consuming (ATP), and therefore recycling of GSSG is important, reducing energy consumption and decreasing the need for available cysteine.
Glutathione is necessary for sperm head swelling, and the impact of glutathione mobilization on further embryonic development is immediate: an increase in the rate of blastocyst formation is observed, with increased cell number per blastocyst formed. This is probably due to the universal role of glutathione in protection against oxidative stress.
Glucose
Sugar metabolism is complex; active transport of hexoses has been shown in mouse embryos, and glucose and lactate are necessary for mouse embryo development in vitro. Hexoses are essential for energy generation during preimplantation development: 1 mole of glucose generates from 30 to 36 moles of ATP. However, the equilibrium between sugars and other metabolic compounds is of paramount importance. It has been suggested that glucose is toxic during in-vitro culture before genomic activation and that glucose and phosphate together may inhibit early embryo development. The mechanisms proposed include induction of glycolysis at the expense of substrate oxidation, through disrupted mitochondrial function. High levels of glucose can lead to excessive free radical formation, but the toxic effect is dependent upon the overall composition of the media; in some systems the negative effect of glucose may be counterbalanced by the presence of a correct amino acid balance, i.e., the presence of sulfur amino acids and derivatives that neutralize reactive oxygen species (ROS). Amino acids and EDTA suppress glycolysis through different combinations, and act in combination to further suppress glycolysis. After genome activation, glucose becomes a key metabolite, required for lipid, amino acid and nucleic acid synthesis. It is also essential for blastocyst hatching.
Lipids
Lipids are essential for very early stages of development: meiotic resumption and oocyte competence depend on lipid beta-oxidation, and lipids are then required immediately after fertilization. Three times more ATP can be generated from one chain of fatty acid (palmitate) than from one molecule of glucose. Mitochondria are involved in the metabolism of lipids, and this requires the presence of carnitine as a catalyst. Lipids can be synthesized (through C-2 condensation reactions), accumulated from the surrounding medium or carried with albumin. Cholesterol synthesis is possible, but slow: there is a rate-limiting step at the level of hMG (3-hydroxy-3-methyl-glutaryl) CoA reductase. If synthesis of cholesterol is experimentally inhibited by chetosterol, the embryos arrest and die.
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