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Chapter 2
Embryonic Development of the Ovary,
Sexual Reproduction and Meiosis
Gerardo H. Vázquez-Nin, María Luisa Escobar, and Olga M. Echeverría
Abstract Sex first evolved in the common ancestor of all eukaryotes some two
billons years ago. Fertilization is the union of two haploid gametes of different
sex; the resulting diploid cell is the zygote. In mammalian species gametes are
derived from precursors termed primordial germ cells (PGCs). Specification of the
germline occurs through: (1) repression of somatic differentiation; (2) reacquisition
of ­potential pluripotency; (3) genome wide epigenetic reprogramming. In several
mammals the maintenance of germ cell linage is not due to a preformed “germ
plasma”, but is induced by environmental signals. In mammals PGCs originate in
extraembryonic region and eventually move to gonadal ridges, located in the medial
part of the urogenital ridge. Observations and experimental evidence demonstrate
that PGCs move actively from the hindgut to the gonadal ridges. However, some
observations suggest that most displacements of these cells may be explained by
global growth movements. In most mammals sex determination is genetically controlled by the presence or absence of Y chromosome and the expression of gene
Sry. However, the ovary differentiation requires in addition the action of a precise
gene cascade. The main process that marks PCGs/oogonia sex differentiation
within fetal ovary is the beginning of meiosis. During the progression of the first
meiotic prophase many germ cells are eliminated, around 70% in mice, rats and
humans. Prenatal development of the follicles is regulated by a complex interplay
of hormones produced by the hypophysis (FSH, LH), and paracrine regulatory factors produced by the oocytes and the surrounding somatic cells. Epigenetic signals
are covalent modifications of histones or DNA that convey a specific “meaning” to
the stretch of chromatin on which they are found. Frequently they induce changes
in chromatin conformation and gene expression. Pluripotent embryonic germ
cells are capable of giving rise to all tissues, thus they must be able to reorganize
their mechanisms of gene regulation to allow multiple lines of cell differentiation.
Besides epigenetic modifications regulate X inactivation/reactivation in the female
G.H. Vázquez-Nin (*), M.L. Escobar, and O.M. Echeverría
Laboratory of Electron Microscopy, Department of Cell Biology, Faculty of Sciences,
National University of Mexico (UNAM), Mexico City, Mexico
e-mail: [email protected]
G.H. Vázquez-Nin et al., Cell Death in Mammalian Ovary,
DOI 10.1007/978-94-007-1134-1_2, © Springer Science+Business Media B.V. 2011
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embryo and mark the paternal and maternal genome of germ cells, making the
expression of a group of genes monoallelic, crucial for embryo development termed
for modification such as “imprinted” genes.
2.1 Evolutionary Advantages of Sexual Reproduction
In all mammalian species, gametes are derived from precursors termed the
­primordial germ cells (PGCs) which migrate into and colonize the gonadal ridges
during embryonic development. PGCs and the cells originated from them, the
oogonia in the female and the spermatogonia in the male, are diploid cells, as any
somatic cell of the body of eukaryotes. This means that they have a pair of each
chromosome formed by one chromatide (monochromatide chromosome) during
interphase and two chromatides (bichromatide chromosome) when the cell is ready
for division. The mature gametes (secondary oocytes or eggs and spermatozoa) are
instead haploid cells since they have only one monochromatide chromosome of
each pair.
Meiosis is the germ cell exclusive cell division producing haploid cells. Meiosis
involves two cell divisions with only one DNA duplication and results in four
­haploid spermatids in the male or one mature egg or ovum and two small cells
called polar bodies in the female.
Fertilization is the union of two haploid gametes of different sex resulting in a
diploid cell termed zygote. Meiosis and fertilization are the two complementary
processes of the sexual reproduction cycle, the reproductive process most favored
by the evolution of eukaryotes.
Sex first evolved in the common ancestor of all eukaryotes some two billion years
ago (Zimmer 2008). To explain the rarity of asexual eukaryotes, even if they can
rapidly spread a beneficial mutation in a population, they can pass mutation only to
their direct offspring. Thus the beneficial mutation will be always inserted in the
same genome. If another organism undergoes a different beneficial mutation in a
different gene, both mutations cannot be combined in the one genome. Sexual reproduction recombines genes, joining beneficial mutation and possibly improving a
population faster than asexual reproduction. Some examples testify the superiority
of sexual reproduction over asexual. Genetically engineered yeast that could only
reproduce sexually and others that could only reproduce asexually were raised on a
near-starvation diet. Sexual yeast was able to adapt faster and as they evolved, their
growth rate increased 94%, while asexual strain increased only 80%. This difference
in growth allowed the sexual yeast to take over the population (Zimmer 2008).
Asexual organisms may pick up slightly deleterious mutations that natural selection
fails to eliminate. In this way slight deleterious mutations may replace undamaged
genes in a population and permanently compromise fitness. Sexual organisms due to
meiotic recombination may interchange a defective gene for a working one and keep
healthy genomes (Zimmer 2008). A long term study (30 years) analyzed the relationship between two snail populations, Potamopyrgus antipodarum, one asexual
2 Embryonic Development of the Ovary, Sexual Reproduction and Meiosis
27
and other sexual, with a parasitic trematode capable of sterilizing them. At the
­beginning the most common strains of asexual snails were resistant to the trematode.
Over the years, however, the snails became progressively vulnerable to an adapting
strain of trematode. Ten years later, the initial strain of asexual snails had almost
disappeared and the population of sexual snails remained relatively stable (Zimmer
2008). In a study of sexual and asexual lineages of the microcrustacean Daphnia
pulex it was found that deleterious amino acid substitutions in mitochondrial ­proteins
are four time faster in asexual lineages than in sexual ones. These results demonstrate that sexual reproduction reduces the deleterious mutational burden in the
mitochondrial genome as well (Paland and Lynch 2006).
2.2 Strategies for PGC Formation
PGCs form outside the gonads during early stages of embryonic development and
enter or are subsequently enclosed into the developing gonads (gonadal ridges)
where they differentiate into primary oocytes in the female (Fig. 2.1) and
­prospermatogonia or gonocytes in the male.
The specification of PGC precursors takes place very early in development in
Chordata, Nematoda and several other Phyla. Specification and determination of the
germline occur through the integration of three key events: repression of the somatic
Fig. 2.1 Primordial germ cells reach the gonadal ridges to colonize the gonad. A close-up is
shown upper the panel
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differentiation program, reacquisition of potential differentiation pluripotency and
ensuing genome-wide epigenetic reprogramming (Yamaji et al. 2008).
Early studies in the mouse embryo showed that germ cell lineage is induced in cells
of the proximal epiblast by the extra-embryonic ectoderm (reviewed in Buehr 1997).
Thanks to very elegant, recent studies, crucial events of such a fascinating process
have been revealed. The bone morphogenetic proteins (BMPs) have been shown to
be central in PGC formation in the mouse. These proteins are members of the
transforming growth factor b (TGFb) superfamily that includes TGFßs and
BMPs, GDFs (Growth Differentiation Factors), activins, inhibins, MIS (Müllerian
Inhibiting Substance), Nodal and Lleftys (Chang et al. 2002). The first indications
of the crucial role of BMPs in PGC formation came from the observation that in
Bmp4 or in Bmp8b knockout mice, PGCs were reduced or absent. Mice double
heterozygous for Bmp4 and Bmp2, a close relative of BMP-4 also have a severe
reduction of PGCs (Ying et al. 2001; Chang et al. 2002). Further investigations
demonstrated that the induction of mouse PGCs actually depends on the synergistic
action of BMP4, BMP8b and BMP2 produced by extraembryonic tissues surrounding the epiblast (Fig. 2.2). Transplantation of epiblast cells at different sites, gave
rise to PGCs only if they were positioned in close proximity to the extraembryonic
ectoderm, indicating that factors produced in this tissue are required for the generation of the PGC precursors (Ying et al. 2001). These factors were identified as
BMP4 and BMP8b, while BMP2 was found to be secreted by the extraembryonic
visceral endoderm (Ying and Zhao 2001; see also the reviews by Chang et al. 2002;
De Felici et al. 2004; Van den Hurk and Zhao 2005).
Although similar PGC specification events are likely to occur within the epiblast
of the human embryo around the end of the second week of development before
gastrulation, the different organization of the mouse and human embryo at this time
Fig. 2.2 Schematic representation of the hypothetic synergistic action of BMP4, BMP8b and
BMP2 produced by extraembryonic tissues surrounding the epiblast to induce PGC precursors in
the human embryo around the beginning of the gatrulation period
2 Embryonic Development of the Ovary, Sexual Reproduction and Meiosis
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makes difficult to identify the mouse equivalent extraembryonic tissues responsible
of the inductive germline specification action in humans. We can speculate that the
mouse extraembryonic ectoderm and visceral endoderm correspond to the roof of
the proamniotic cavity and hypoblast (primitive endoderm) of the human embryo,
respectively. As in the mouse, it is likely that from the epiblast, human PGC precursors around the beginning of the third week move together with the extraembryonic
mesoderm cells through the forming primitive streak (Fig. 2.2) toward a region of
the wall of the yolk sac adjacent to allantois where they are determined and have
been identified around 20th day in very old studies (reviewed in Baker and
Eastwood 1983).
Since activation of intracellular signaling by BMPs occurs through SMAD
­proteins, it was not surprising to find that Smad5 and Smad1 knockout mice have
defects in PGC development resembling that caused by the ablation of the BMP
genes (Chang et al. 2002).
These findings demonstrate that in mouse and other mammals, the maintenance
of germ cell lineage is not due to the inheritance of a preformed “germ plasm”, but is
induced by environmental signals acting on pluripotent somatic cells (Saitou et al. 2002).
In other species, however, the germlineage is specified by a different strategy. For
example, in the nematode Ascaris the germ cell linage is segregated during the first
few cleavage divisions of the embryo in cells retaining their chromatin complement
complete, while the somatic lineages suffered chromatin diminution for the loss of
terminal regions of the chromosomes (see review by McLaren 2003). Similar
mechanisms of intrinsic germline specification occur in Xenopus, Frog, Drosophila,
Caenorhabditis elegans. In such species, a region of the cytoplasm of unfertilized
egg near the vegetal pole contains contains aggregates of mitochondria, proteins and
RNA. Cells that during the first mitotic division receive this special cytoplasm
region, the “germ plasm” become PGCs. (see review by McLaren 2003).
2.3 Molecular Mechanisms of PGC Specification in Mouse
To determine the molecular mechanisms of the PGC specification, Saitou and
­collaborators (2002) performed genetic screens between PGCs and their somatic
neighbors that share a common ancestry. The expression of Fragilis, an interferoninducible transmembrane protein, marks the beginning of germ cell competence.
Using single-cell gene expression profiles, these authors demonstrated that the
cells, at 7.25 days post coitum (dpc), show the highest expression of Fragilis, subsequently expressed Stella. The function of Stella gene product is uncertain but it
has domains characteristic of protein involved in RNA splicing. Actually, Stella
represses homeobox genes in nascent germ cells, and as such maintains the pluripotency of PGCs during their migration to the gonadal ridge. The induction of
Fragilis by extra-embryonic tissues is dependent on BMP4 (Saitou et al. 2002).
Wnt3 (a secreted signaling protein transmitted via beta-catenin), provides epiblast
cells with the ability to respond to BMP signals (Ohinata et al. 2009). Going further
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back, at 6.25 dpc, about six cells within the Fragilis-positive cells, at the posterior
side of the embryo, begin to show expression of Blimp1/Prdm1and shortly after of
Prdm14. The restricted expression of these two transcription factors in the PGC
precursors is likely due to the inhibition of BMP4-2 action in the anterior region by
antagonists produced by the anterior visceral endoderm (AVE) (Ohinata et al. 2009).
Experiments tracing genetic lineage demonstrate that all Blimp1 positive cells are
the lineage restricted PGC precursors (Ohinata et al. 2005; Vincent et al. 2005).
The complex Blimp1/Prmt5 arginine methyltransferase mediates a change in the
methylation of arginine 3 on histone H2A and H4 tails between 8.5 and 10.5 dpc.
This change, accompanied by the Prdm14 action, causes other epigenetic changes
necessary for repression of somatic program and ensures that PGCs retain or reacquire a pluripotent character. The Blimp1-positive PGC precursors while proliferating and moving into the extraembryonic mesoderm (exM) re-express key
pluripotency-associated genes such as Sox2, Nanog and Oct-4. This latter belongs
to the class V of the POU domain transcription factor family and has been the first
transcription factor demonstrated to be lineage restricted in the developing PGCs
(for a review, see Pesce et al. 1998). Oct-4, Sox2 and Nanog are transcription
­factors all essential for maintaining the pluripotent stem cell phenotype.
Around 7. 25–7.5 dpc, the PGC precursors in the exM at the basis of allantois are
determined in a cluster of about 40–50 PGCs by unknown factors, but likely still
including BMP-4. The adhesion molecule, E-cadherin is involved in clustering such
PGCs and appears to be crucial for these precursors to be definitively allocated to the
germ cell lineage (Okamura et al. 2003). Determined PGCs, besides the classical
PGC markers tissue not specific alkaline phosphatase (Tnap) and stage-specificantigen-1 (Ssea-1), express Kit receptor and germline-specific genes such as Nanos3
and Dead endI. What activates germline genes in PGCs? Paradoxically, recent evidence from mouse and C. elegans suggests that the same factors responsible for
transcriptional repression may also be involved in such activation. Single-cell transcriptome profiling has shown that Blimp1 is required, not only for virtually all transcriptional repression, but also for the activation of ~50% of the genes that are
upregulated in mouse PGCs. Genes that require Blimp1 for activation include those
that show the highest specificity for PGC expression (e.g. Nanos3) (Kurimoto et al.
2008). The conclusion is that Blimp1 activity is required, directly or indirectly, to initiate part of the germline-specific transcriptional program. Being a Blimp1-expressing
PGC precursors is not sufficient, however, to undergo specification to PGCs. It is
likely that a second signal is needed locally to trigger specification, with another
Blimp1-independent pathway also functioning in parallel (Kurimoto et al. 2008).
This second pathway appears to be controlled by Prdm14 (Yamaji et al. 2008).
Prdm14 expression in PGCs initially occurs independently of Blimp1, but becomes
dependent on Blimp1 by E7.5. In the absence of Prdm14, presumptive PGCs fail to
downregulate GLP (G9a-related protein), the H3K9 methyltransferase, and
­maintain H3K9me2 and do not upregulate H3K27me3. Prdm14-deficient PGCs
also do not activate the pluripotency-related gene Sox2. How Prdm14 and Blimp1
contribute to both the upregulation of certain genes and the downregulation of
­others is not known.
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2.4 PGC Migration
Since the nineteenth century, it was established that in mammals PGCs originate in
extraembryonic region, and eventually move to the gonadal ridges (Nussbaum 1880
quoted by Merchant and Alvarez 1986). During this pregonadic phase PGCs can be
identified by morphological criteria and surface markers, such as TNAP and SSEA-1,
and the expression of genes typical of pluripotent cells (e.g. Oct-4, Sox2, Nanog)
(Oktem and Oktay 2008). From the site of their determination, the wall of the yolk
sac adjacent to the allantois, PGCs reenter into the embryo proper following the
morphogenic movements incorporating the proximal part of the yolk sac into the
hind gut. It is likely that at this stage PGCs acquire a motility phenotype and consequently after invading the hind gut endoderm move throughout the dorsal mesentery towards the gonadal ridges formed in the celomic epithelium.
Ultrastructural observations favored the idea of preexisting pathways from the
hind gut basal lamina to the celomic epithelium. Such pathways are formed by
elongated mesenchymal cells and oriented elements of the extracellular matrix as
collagen fibrils and fribronectin (Merchant and Alvarez 1986). Studies in vivo and
in vitro demonstrated that at this stages mouse and human PGCs are able to emit
filopodia (Fig. 2.3) by which they attach to different structures and retract pulling
Fig. 2.3 Migrating primordial germ cells present filopodia. Migrating cells advance in gradients
of increasing chemoattractant concentration leading to polarized cell protrusions originating the
filopodia. The chemorepellent are avoided for the migrating cells
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the cell to the point of attachment (Albrecht-Buechler 1976; Gustafson and Wolpert
1963). Therefore, locomotion may be atypical ameboidal (Zamboni and Merchant
1973; Merchant and Alvarez 1986). Taking into account these data, Merchant and
Alvarez (1986) proposed: (1) the presence of short pathways containing fibronectin
between the hind-gut and the celomic epithelium; (2) that fibronectin is required for
PGC locomotion and (3) that mammalian PGCs emit filopodia which establish
contact with surrounding substrata. Contact guidance migration of PGC might be
mediated by KIT receptor expressed on the surface of these cells and its ligand, Kit
ligand (also called Stem cell factor), which is expressed in somatic cells along the
migratory pathway (Fleischman 1993; van den Hurk and Zhao 2005). Kit ligand
also stimulates PGC migration, survival and proliferation in vitro (Dolci et al. 1993).
Molyneaux et al. (2001) studied systematically the displacement of the PGCs in the
mouse embryo, concluding that during some stages of the migration between the
hind gut and the gonadal ridges PGCs move actively and directional to their final
destination. More recently, in a time-lapse comparative analysis of the PGC migration between Drosophila, Zebrafish and mouse, these authors defined six distinct
stages of mouse PGC behavior: (1) invasion of the endoderm; (2) ­passive or active
migration into hindgut; (3) random migration within the hindgut; (4) migration from
the gut to the gonadal ridges; (5) clustering at the ridges; (6) cell death within midline structures (Molyneaux and Wylie 2004). At stage 3 the PGCs are located in the
hindgut epithelium moving actively around the epithelial cells. The epithelial cells
express the Ca++ dependent adhesion molecule E-cadherin; whereas PGC do not.
The lack of strong adhesive interactions allow PGCs to move freely within the gut.
PGCs upregulate E-cadherin expression upon leaving the gut (Molyneaux and
Wylie 2004; Di Carlo and De Felici 2000). Then PGCs exit from the gut, divide in
two streams and migrate to the developing gonadal ridges. In organ culture of transverse slices of mice embryos, the addition of BMP4 elevates the PGC number. The
addition of Noggin, a BMP-antagonist, reduced the PGC number and slowed and
randomize their movements, resulting in failure of PGCs to colonize gonadal ridges
(Dudley et al. 2007). The action of BMPs is mediated by the BMP-specific Smads
(Smad1, Smad5 and Smad8). Immunolocalization of Smads1/5/8 demonstrates that
migratory PGCs do not express detectable levels of these proteins. Instead the
somatic cells of the gonadal ridges exhibit an intense Smads1/5/8 immunolabeling,
thus suggesting that BMPs act on these cells (Dudley et al. 2007). Several growth
factors contribute to enlarge the PGCs population. The migratory PGCs of mouse
embryos express subunits of interleukin-2 receptor. The addition of Interleukine-2
to PGS in culture increased their number by a mitogenic effect. These data are
indicative of a paracrine effect of Interleukine-2 on PGC proliferation (Eguizabal
et al. 2007). When mouse PGCs were co-cultured with gonadal ridges they increased
in number and moved towards the gonadal ridges. These effects were present at long
distances and were absent when PGCs were co-cultured with other embryonic
organs. Gonadal ridge conditioned medium attracts PGCs and TGFß may mimic
these effects (Godin et al. 1990;Godin and Wylie 1991). These results strongly suggest that the cells of gonadal ridges secrete chemo-attractants (van den Hurk and
Zhao 2005). More recently, some evidence that KL itself exerts chemiotactic effect
on mouse PGCs in vitro was reported (Farini et al. 2007).
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In spite of the numerous observations and experimental evidence demonstrating
that PGCs move actively, some observations in mouse and human embryos suggest
that most of the displacements of these cells can be explained by the global growth
movements of the embryo and that possibly no active migration exists at all
(Freeman 2003). The same author concludes that the displacement the PGCs
in vivo are unrelated to the “artefactual” movements of explanted germ cells.
2.5 Formation of the Ovary
The gonads develop in the medial part of the urogenital ridge. The urogenital ridges
are longitudinal thickenings located at both sides of the dorsal mesentery. The
gonadal ridge is unique among all organ primordia because of its bipotential nature.
A testis or an ovary can develop from a gonadal primordium. This first appears
as thickenings of the celomic epithelium. As the epithelium invades the connective
tissue the genital ridges protrude to the celomic cavity forming the gonadal ridge.
This proliferation of the epithelium results in an internal epithelial mass and
the stratified superficial epithelium of the gonadal ridge.
In most mammals sex determination is genetically controlled by the presence
or absence of a Y chromosome. The expression of the Y linked gene Sry in mice
starts the rapid differentiation of testis specific cell types. In the absence of Sry
even an XY mouse develops ovaries (Brennan and Capel 2004). The formation of
other sexual organs and tissues is largely a consequence of the sex differentiation
of the gonads and their hormone production. On the other hand, the targets of the
molecular mechanisms of sex differentiation are the somatic cells of the gonads
and not the germ cells. PGCs are actually sexually bipotent and differentiate in
oogonia/oocytes or prospermatogonia under the influence of the somatic environment of the ovary and testis, respectively, and independently by their sex chromosomes (Fig. 2.4).
Recent results indicate, however, that ovary differentiation does not simply
occur for default in the absence of Sry but requires the action of a precise gene
cascade. Actually, the “canonical” ß-catenin signaling pathway represents a central
piece of the mechanisms controlling gonad differentiation into ovary. This signaling is initiated by WNT ligand binding to Frizzled and LRP5/6 co-receptors,
­leading to the inhibition of the ß-catenin destruction complex, containing glycogen
synthase kinase-3-beta (GSK-3-beta), the adenomatous polyposis coli protein
(APC), and the scaffolding protein axin, among others. This complex phosphorylates ß-catenin and targets it for degradation by the proteasome. Its inhibition leads
to ß-catenin stabilization and entry to the nucleus. Nuclear ß-catenin binding to
T-cell factor (TCF)/ lymphoid enhancer factors (LEF) leads to the activation of
target gene expression. In the absence of ß-catenin, TCF/LEF proteins act as a
transcriptional repressor. This ability to switch between repressor and activator
states, probably tighten control over gene expression (see review by Tevosian and
Manuylov 2008). Several Wnt family members are expressed in the developing
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Fig. 2.4 Schematic diagram showing that the targets of the molecular mechanisms of sex
­differentiation are the somatic cells of the gonads and not the germ cells
mammalian gonads. Initially there is a low level of expression of Wnt4 in both XX
and XY gonadal primordial. This expression is down regulated in differentiating
male gonad while it is increased the female. In mice, Wnt4-null females are
­masculinized as by the absence of Müllerian ducts, a dramatic decrease in the number of developing oocytes, production of ectopic steroids as testosterone and formation of male specific blood vessels. Markers of early ovarian development as
Follistatin and Bmp2 are not expressed in the absence of Wnt4. The effects of the
increase of Wnt4 dosage during embryogenesis in mice is relatively mild, mostly
confined to the abnormal development of celomic blood vessels in the male. In
summary, the absence of Wnt4 expression is insufficient by itself to program the
testis differentiation of somatic cells. However, Wnt4 controls several elements of
sexual development (see review by Tevosian and Manuylov 2008). Rspo is another
gene family with a role in female sexual development which also targets the
ß-catenin transduction pathway. A recessive mutation of the gene encoding RSPO1
results in a complete female to male sex reversal in humans. The Rspo1 mutation
(XX Rspo1-/-) in mice results in a phenotypical female. However, the gonads
undergo incomplete sex reversal and the masculinization occurs postnatally. The
conclusion seems to be that Rsp1 plays a positive role in mammalian ovary development, but the genetic mechanism of the late sex reversal remains to be determined. It is well known that activating Sry or Sox9 in the female will override the
ovarian pathway and activate the male program in a XX gonad. A human XY
patient carrying a chromosome duplication which included the WNT4 and RSPO1
genes exhibited sex reversion. This suggested that the up-regulation of both of these
2 Embryonic Development of the Ovary, Sexual Reproduction and Meiosis
35
ligands in mice may be enough to antagonize male development. Because both
WNT4 and RSPO1 signaling converge in activating canonical ß-catenin pathway,
the researchers produced an undegradable form of ß-catenin in somatic cells of the
developing ovary. This was sufficient to block the male pathway in the XY gonads.
The stabilization of ß-catenin blocked male-specific Sox9 and Amh expression,
while genes normally found in the ovarian somatic cells as Foxl2, Bmp2, Wnt4 and
Fst were increased. Testis cords did not form in these XY mutants (see review by
Tevosian and Manuylov 2008).
In summary we can conclude that sexual differentiation of the gonads in
­mammals is driven by two opposing antagonistic activities of the transcriptional
regulatory proteins: SOX9 in males and ß-catenin and T-cell factor (TCF)/lymphoid
enhancer factor (LEF) in females (see review by Tevosian and Manuylov 2008).
2.6 Germ Cell Sex
For the colonization and development of the PGCs into the gonadal ridges the
expression of a large number of genes is needed (van den Hurk and Zhao 2005).
The proliferation of somatic and germ cells leads to the enlargement of the gonadal
ridge. In the developing ovary, PGCs (in some species now called oogonia) interact with epithelial somatic cells of the ovary and give rise to ovarian or ovigerous
cords, which are continuous with the surface epithelium. These cords are more
clearly defined in some species that in others (Pepling 2006). Numerous epithelial
cells surrounding the PGCs in the ovigerous cords originate in the mesonephros.
These epithelial structures are delimited by a basal lamina and surrounded by
mesenchymal cells (Guigon and Magre 2006). Other studies support the nonmesonephric origin of gonadal somatic cells. They are mainly based in the demonstration that mesonephric agenesis does not block the formation of the gonad.
The cells in the origin of the mesonephros, the adrenal gland and the gonad correspond to a multivalent urogenital mesenchyme (Merchant-Larios 1985). Groups
of oogonia that are surrounded by a layer of epithelial cells (granulosa cells) form
cysts. In human and other mammals oogonia are connected by cytoplasmic bridges
giving rise to a clonal proliferation (Pepling 2006). Mitotic activity of oogonia
determine the maximum number germ cells in the life span, that is six to seven
million at 20 weeks of gestation in the human (Oktem and Oktay 2008). Due to a
high rate of programmed cell death, a girl has one million oocytes at birth and
three to four hundred thousand at puberty. Of these only 300–400 will ovulate
until menopause (Oktem and Oktay 2008).
The main process that marks PGCs/oogonia sex differentiation within the fetal
ovary is the beginning of meiosis. PGCs/oogonia entering into meiosis are termed
primary oocytes. These pass through the stages of prophase I (leptotene, zygotene,
pachytene and diplotene), and around the time of birth become arrested at dictyate
stage, which is a pseudo-interphase stage corresponding to the last G2 stage of meiotic
process. In contrast, the spermatogonia are not engaged in meiotic process until puberty.
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The debate whether the PGCs/oogonia enter into meiotic S phase ­autonomously
following internal signals (Zamboni and Upadhyay 1998; McLaren and Southee
1997) or induced by external signals is still undergoing (reviewed in Bullejos and
Koopman 2004). In any case, as reported in the previous section, entering into meiosis does not depend on sex chromosomes. Recent studies demonstrate that retinoic
acid (RA), a derivate of vitamin A, induces oogonia to enter in meiosis during fetal
life in the mouse ovary and that RA degradation in the fetal testis ­prevents spermatogonia to do the same at the same time (see review by Bowles and Koopman
2007). Stimulated by retinoic acid, gene 8 (Stra8) is involved in the initiation of the
meiotic process. For the decision, it is required to enter in the S phase preceding
meiosis (Baltus et al. 2006). Germ cells do not initiate meiosis in Stra8-null mice, so
RA must trigger meiosis inducing the cytoplasmic protein STRA8, which is never
expressed in gonadal somatic cells, possibly due to differences in the epigenetic
configuration of the regulatory regions of the gene. Retinoic acid ­produced in the
mesonephros, causes germ cell in the ovary to enter meiosis (Bowles et al. 2006),
and is also the key to sex-specific timing of meiotic process (Bowles and Koopman
2007). It is likely that besides RA availability entering into meiosis in female PGCs
or meiotic prevention in male PGCs require additional molecular control that is
being to be identified. For example, epigenetic modification of chromatin in female
PGCs and the action of FGF-9 in male PGCs are involved in favoring or preventing
meiosis, respectively (Wang and Tilly 2010; Barrios et al. 2010).
As reported above, oocytes ring into the last stage of prophase I become
arrested in dictyate. The molecular mechanisms of such arrest, necessary for
allowing the progressive recruitment of group of oocytes into the growing phase
after birth, are unknown in mammals but are likely to involve players of cell cycle
control such as cyclins, CDKs and CDKI (Western et al. 2008; Spiller et al. 2009;
Miles et al. 2010).
When the meiosis is arrested in diakinesis at the end of the prophase of the first
meiotic division, a prolonged pseudo-interphase takes place. Around and early after
birth, ovigerous cords brake down in follicles, the oocytes are progressively
­surrounded by a monolayer of flat epithelial cells, now called granulosa or follicular cells and a basal lamina, giving rise to the primordial follicles. This process
begins in the central region of the ovary and proceeds to the periphery. These follicles are surrounded by a microvascular network of capillaries juxtaposed to basal
lamina (Sato et al. 2006). The development of primordial follicles seems to be
related to meiotic dyctiate stage block (Borum 1961). Experimental studies on the
relation of a protein of the synaptonemal complex (SCP1), which is typical of
pachytene stage and the formation of primordial follicles, demonstrate that the
SCP1 deficient ovaries exhibited an increased number of newly formed follicles.
These findings suggest that the completion of pachytene stage or probably an artificial finalization of meiotic prophase I endow the oocytes with the ability to
orchestrate the follicular assembly (Paredes et al. 2005). Spontaneous primordial
follicle development was completely inhibited by a Kit antibody that blocks
­kit-ligand/stem cell factor actions. Experiments in vitro with rat ovaries demonstrated that kit-ligand is necessary and sufficient to induce primordial follicle
2 Embryonic Development of the Ovary, Sexual Reproduction and Meiosis
37
d­ evelopment. On the other hand, gonadotropins (FSH and hCG) do not induce
follicle ­development (Parrot and Skinner 1999).
During the progression of the first meiotic prophase and follicle formation many
germ cells are eliminated, around 70% in mice, rats and humans. The reasons for
this constitutive germ cell death are not clearly established and will be discussed in
other chapters of this book.
It is important to note that germ cells are necessary for the correct formation of
the ovary. However, if PGCs are eliminated by busulphan treatment, a drug that
selectively destroys them, (Merchant 1975) or by the pleiotropic mutation W/Wv
(Merchant-Larios and Centeno 1981), in the rat and in the mouse, somatic cells
undergo proliferation and form an undifferentiated gonad in both sexes. That is, a
gonad consisting in packed cells that gradually become organized into two different
tissues separated by a basal lamina: epithelial layers and/or epithelial cords and the
stromal tissue (Merchant 1975). Similar results have been obtained in Amphibia
(Bounoure 1950; Padoa 1964) and in chicks (Willier 1950). Fragmentation of the
cords to form follicles takes place only when germ cells act as differentiating centers (Merchant 1975). Some PGCs survive in the gonad of mutant W/Wv mice and
are able to transform in normal oocytes and organize follicles capable of inducing
steroidogenic tissue (Merchant-Larios and Centeno 1981).
2.7 Endocrine and Paracrine Regulation of Folliculogenesis
In order to understand the prenatal endocrine relation of the developing ovary it is
necessary take into account the developmental relationships between the
­hypothalamus, the hypophysis and the gonad. Anterior pituitary is formed by an
evagination of the oral ectoderm and the posterior lobe is formed by a ventral prolongation of the diencephalon. Autocrine and paracrine signaling are involved in
the initial development of the gland. This system differentiates and acquires capability for endocrine function very early. In humans, luteinizing hormone and follicle
stimulating hormone are produced in the pituitary as early as the fifth week of gestation. Gonadotropin releasing hormone and other releasing hormones are present
in the hypothalamus at about the same time. The ensemble can function as a whole
after the differentiation of the portal vessels between the hypothalamus and pituitary (Cummings and Kavlock 2004). When the gonads begin to produce hormones,
interactions take place between the gonads and the hypothalamic-pituitary axis.
In the female human fetus estrogen is present by the 10th–14th week, but the hormone
peaks at about the 20th week, following increments in hypothalamic gonadotropin
releasing hormone and pituitary gonadotropin concentrations. Near birth, the hypothalami of the female acquire the capability to respond to rising estrogen levels and
the triggering of the surge of LH that is a requisite for ovulation. However, these
events will not occur until puberty (Cummings and Kavlock 2004).
The development of the follicles is regulated by a complex interplay of hormones
produced by the hypophysis as follicle-stimulant hormone (FSH) and luteinizing
38
G.H. Vázquez-Nin et al.
Fig. 2.5 Folliculogenesis process. The preantral follicles are independent of gonadotropin. The
development of the follicles is regulated by FSH and LH, and paracrine regulatory factors
­produced by the oocytes and the surrounding somatic cells
hormone (LH), and paracrine regulatory factors produced by the oocytes and the
surrounding somatic cells. The latter are especially crucial for the formation of the
primordial follicle population around and early after birth and likely for the selection
of follicles from these populations to be driven through the subsequent maturation
stages. Hormones begin to play a role from the primary follicle stage throughout the
secondary, preantral and antral follicle stages up to ovulation (Fig. 2.5).
FSH up regulates a number of genes in granulosa cells, such as cyclin D2
involved in proliferation, aromatase enzyme required for estrogen production, and
LH receptor (Anttonen 2005). FSH receptor expression is restricted to granulosa
cells, whereas LH receptor is expressed both in granulosa and theca cells. LH
stimulates androgen synthesis in theca cells and FSH induces estrogen conversion
from androgen, by means of aromatase enzyme, in granulosa cells (Gómez et al. 2001).
The defects in folliculogenesis differ between FSH receptor- and LH receptordeficient mice. Both have no mature, preovulatory follicles or luteal glands, but in
FSH receptor-null mice follicular growth arrests at the preantral stage, whereas in
LH receptor-null mice the follicles arrest at the antral stage. These findings demonstrate the need for FSH action at the preantral stage and beyond, and both FSH and
LH actions for the follicular maturation and ovulation stages (Anttonen 2005).
Oocytes and somatic cells produce several paracrine factors, modifying gonadotropin actions. Oocyte supplies specific morphogens that preserves phenotype and
gene expression in cumulus cells. On the other hand, the somatic cells provide the
oocytes with nutrients and growth factors during folliculogenesis (Anttonen 2005).
Members of the TGF-ß superfamily have essential roles in these paracrine interplays. Growth and differentiating factor 9 (GDF9), bone marrow-morphogenic
2 Embryonic Development of the Ovary, Sexual Reproduction and Meiosis
39
protein 15 (BMP-15), and BMP6 are secreted by the oocyte, while activins, ­inhibins
and anti-Müllerian hormone (AMH) are expressed and secreted by somatic cells.
AMH, in addition to its role in the differentiation of male reproductive tract, is a
granulosa cell product capable of suppressing follicular growth induced by FSH
and epidermal growth factor. AMH-null mice have an increased number of growing
follicles, resulting in loss of the primordial follicles at a premature age (Anttonen 2005).
For a general physiology text book consult Peterson (2007).
2.8 The Meiotic Division
The meiosis is composed of two cell divisions preceded by the only S phase of this
process (Fig. 2.6). The result is four cells with half the number of chromosomes of
the species and only one DNA double helix per chromosome. The haploid cells are
the mature gametes, the oocytes and the spermatocytes. The first division is composed of an S phase followed by G2 or preleptotene stage, a long prophase compose
of several stages: leptotene, zygotene, pachytene, diplotene, and diakinesis
(dictyate). Several processes specific to meiosis take place during this prophase
Fig. 2.6 Schematic drawing of meiosis process (kindly provided by Abrahan HernándezHernández)
40
G.H. Vázquez-Nin et al.
such as homologous recognition, alignment of homologous chromosomes, pairing
of homologues by means of a special structure, the synaptonemal complex, and
chiasm formation. The oocyte remains in diakinesis until ovulation when it
­proceeds to the metaphase in which the homologous chromosomes are still associated by the chiasmata, and then to the anaphase of the first division, in which each
member of homologous chromosome pairs migrates to a different pole of the
­spindle. Thus the diploid number of chromosomes is divided between the two
­products of the first telophase. The meiotic spindle is located at the periphery of the
cytoplasm, near the cell membrane and it specifies the position of the cleavage furrow, thus the two new cells formed by the telophase are very different in size. The
larger ­daughter cell corresponds to the secondary oocyte and the smaller is the first
polar body. Each has the haploid number of chromosomes formed by two DNA
double helices. The second meiotic division is when the sister chromatids of each
chromosome separate and migrate to a different cell like in a mitotic division. In this
cytokinesis, as in the first one, the cleavage furrow separates a small polar body and
a large oocyte. The second meiotic division takes place only after fertilization.
The biological role of meiosis is the production of gametes (haploid cells),
which are cells with only one member of each pair of chromosomes formed by one
DNA double helix, frequently formed by paternal and maternal segments. During
meiotic process, genetic recombination takes place by two different processes:
random distribution of maternal and paternal homologues, in the first anaphase, and
recombination. The maternal and paternal homologous forming each bivalent chromosome distribute at random at the first metaphase, so each secondary oocyte may
incorporate a different mixture of paternal and maternal chromosomes. The recombination is the exchange of segments of chromatid between homologous, maternal
and paternal chromosomes, when they are paired at the pachytene stage of the first
meiotic prophase. Each exchange is visualized as a connection called a chiasma
between the chromosomes. The chiasmata are easily seen under a light or electron
microscope in post-pachytene stages of meiotic prophase I.
The ultrastructural and molecular aspects of meiosis are better studied in
­spermatocytes than in oocytes, but most processes taking place in the prophase of
the first meiotic division are similar in both sexes. The phase G2 of the last interphase of the spermatogonia is characterized by small lampbrush structures of
extended chromatin formed by an axis from which irradiates the loops. These
micro-lampbrush chromosomes are not found in interphase nuclei of somatic cells.
They persist during leptotene and zygotene stage. Immunolocalization and in situ
localization DNA and RNAs using electron microscope demonstrate that these
lampbrush structures are formed by extended chromatin transcribed mainly or
exclusively by RNA polymerase II and that the product of transcription is still
associated to the chromatin; they contain introns and ribonucleoproteic particles
associated to splicing are scarce. The results of quantitative autoradiography after
[3H]uridine labeling demonstrate that transcription of the extended chromatin of the
micro-lampbrush is intense but the export of RNA to the cytoplasm is very slow.
Then the newly synthesized RNA is probably involved in a nuclear function taking
place within these cells: the process of homology searching and recognition,
2 Embryonic Development of the Ovary, Sexual Reproduction and Meiosis
41
p­ revious to pairing (Vázquez-Nin et al. 2003). The searching of homologies is
facilitated by a much conserved mechanism among eukaryotes, the bouquet. The
bouquet involves the association of the telomeres of all chromosomes to the nuclear
envelope and the subsequent displacement to a small zone of the envelope near the
centrosome, a microtubule organizing center in the cytoplasm. This mechanism
facilitates finding homologue regions of homologous chromosomes. The final pairing of homologous chromosomes takes place with the formation of another structure extremely conserved by evolution, the synaptonemal complex. This complex
is formed by the parallel alignment of corresponding regions of homologous chromosomes from telomere to telomere, and is typical of the pachytene stage. The
synaptonemal complex is formed by two lateral arms and a central space devoid of
DNA except in the recombination nodules. The lateral arms are formed by DNA,
RNA and proteins. Two filaments of chromatin formed by two DNA double helixes
each are folded inside the lateral arms. These filaments are continuous with the long
loops surrounding the complex. RNA of unknown origin is associated with the
DNA filaments (Vázquez-Nin and Echeverría 1976; Ortiz et al. 2009). The complex
is surrounded by long loops of extended chromatin associated to the lateral arms.
Each lateral arm and the loops associated with it correspond to one of the paired
homologous chromosomes. The DNA of the lateral arms of the synaptonemal complex is composed of non-coding repetitive sequences (Hernández-Hernández et al.
2008). For a recent review of epigenetic aspects of the synaptonemal complex and
its formation (Hernández-Hernández et al. 2009).
The alteration of the structural maturation of the central element of the
­synaptonemal complex abolishes the formation of crossovers, preventing recombination (Hamer et al. 2008). Many genes are involved in the process of recombination. Some of them, such as Spo11, are involved in double strand breaks and are
conserved between yeast and mammal. In mammals, yeast and higher plants,
recombination takes place more frequently in 1–2 kb long regions of the genome
called hot spots (Parvanov et al. 2009). The frequency of recombination is low
outside of hotspots. Recombination hot spots in humans are not associated with
DNase I or microccocal nuclease-hypersensitive sites (Fan and Petes 1996); they
are more influenced by base composition (DNA motif), coding context and DNA
repeats (Myers et al. 2006). In human genome both repeat DNA and exonic
sequences are negatively associated with recombination; indeed recombination
rates reach a peak at distances of 10–50 kb from genes (Myers et al. 2006). On the
contrary, regions of high GC content are strongly associated with hotspots, also the
seven bases nucleotide motive CCTCCCT is five times enriched in the hot spots
of the THE1A and THE1B retrotransposons. Mutation of one base of this motive
strongly reduces the hotspot activity (Myers et al. 2006); protein-DNA complexes
are essential for hotspot activity. The ade6-M26 hotspot (chromosomal region) of
fission yeast is the most extensively characterized and best understood meiotic
hotspot in eukaryotes (Gao et al. 2009). Atf1-Pcr1 protein heterodimer associates
with ade6-M26 hotspot. The homologous recombination –activation domain
resides exclusively in Atf1, which is a member of ATF/CREB family of transcription factors. The Atf1-Pcr1-M26 protein-DNA complex promotes locally the
42
G.H. Vázquez-Nin et al.
i­ nitiation of recombination from double strand breaks catalyzed by Rec12 (Spo11)
recombinase, which is a conserved protein of the meiotic recombination machinery.
The recombination-promoting domain resides in Atf1 protein and the hotspot
activation requires the phosphorylation of ATF1 by Spc1, a protein kinase of the
p38-family. The regulation of basal meiotic recombination is independent of this
phosphorylation, whose function is to correct position of Atf-Pcr1 complex (Gao
et al. 2009). In Saccharomyces cerevisiae double strand breaks occur within a few
hundred base pairs of the transcriptional sites. However, only some hotspots are
activated by transcription factors. The well characterized hotspot located within
the promoter of the gene HIS4 is dependent on several transcription factors and
ablation of the transcription factor binding sites abolishes recombination (Borts
2009). The study of a hot spot in the mouse genome, called Psmb9, showed that
its activity is induced by a locus named Dsbc1, for double strand break control 1.
This locus is located within a 6.7-Mb region on chromosome 17. Dsbc1 influences
double strand breaks and crossovers, not only at Psmb9, but in several other
regions of the chromosome 17. Dsbc1 influences the crossover distribution on
chromosomes 15 and 18, and even the crossover activity in the Hlx1 hotspot in
chromosome 1 (Grey et al. 2009). Recently it was found that the recombination
activity of several hotspots on mouse chromosome 1 can be either activated or
suppressed by the CAST allele(s) of distant trans-acting loci and that the activation
of several hotspots is controlled by the locus Rcr1 that exert its action at the initiation of recombination (Parvanov et al. 2009).
These interesting results on the positioning, the molecular mechanism and the
regulation of meiotic recombination are probably just the beginning of a long line
of future research.
Failure to repair DNA damage caused by recombination triggers meiotic checkpoints and destruction of the germ cells by apoptosis can take place. However, there
is an important sexual dimorphism; checkpoint requirements are much less stringent in females than in males. This provides an explanation for the ten-fold higher
number of meiotic errors in females compared with males (Cohen et al. 2006).
2.9 Epigenetic Reprogramming of Germ Cells
Epigenetic signals are covalent modifications of histones or DNA that convey a
specific “meaning” to the stretch of chromatin on which they are found. Frequently
they induce changes in chromatin conformation and gene expression.
Pluripotent embryonic germ cells are capable of giving rise to all tissues derived
from the three embryonic primary epithelial layers: ectoderm, mesoderm and endoderm. Thus they must be able to reorganize their mechanisms of gene regulation to
allow multiple lines of cell differentiation.
Besides determining genome plasticity, epigenetic modifications regulate X
inactivation/reactivation in the female embryo and mark the paternal and maternal
genome of germ cells during gametogenesis, making the expression of a group of
2 Embryonic Development of the Ovary, Sexual Reproduction and Meiosis
43
genes monoallelic, crucial for embryo development termed for modification such
as “imprinted” genes.
As reported above, BLIMP1 (PRDM1), a transcriptional repressor, plays a
­critical role in the specification of PGCs in mice. BLIMP1 interacts with PRMT5,
an arginine-specific methyltransferase, which mediates symmetrical dimethylation
of arginine 3 in histone H2A and/or H4 tails. At E11.5, BLIMP1-PRMT5 translocates from the nucleus to the cytoplasm, resulting in the loss of H2a/H4 R3 methylation, thus determining an extensive epigenetic reprogramming of germ cells. At
this stage, Dhx38, a putative target of the BLIMP1-PRMT5 complex, is also
upregulated. The expression of Dhx38 is seen in pluripotent embryonic germ cells
that are derived from PGCs when Blimp1 expression is lost. These observations
demonstrate that in mice Blimp1 is involved in posttranscriptional regulatory
actions mediated by methylation-demethylation of histone H2A and/or H4 arginine
3 (Ancelin et al. 2006).
Postranscriptional modifications of the histones may be reprogrammed more
easily than DNA methylation, which is a very stable modification that may be heritable. In mammals methylation of the DNA occurs almost exclusively on CpG
dinucleotide, without preference for sequence context surrounding this target.
About 60% of human genes are associated with CpG islands, of which the great
majority are unmethylated at all stages of development, in all types of tissues (see
review by Bird 2002). It is important to note that the DNA methylation does not
silence active promoters, but affects genes already silenced, making the turn off
permanently. DNA methylation is associated with a stable repression of transcription, frequently accompanied by conformational changes of the repressed loci.
Examples of the modifications due to DNA methylation are monoallelic expression
of some imprinted genes, repression of transcription of retrotransposons and inactivation of one X chromosome, and other examples quoted by Bird (2002). In all
cases there is a compaction of the repressed chromatin and the modifications are for
the lifespan (Lees-Murdock and Walsh 2008). DNA methylation is heritable
through cell division by copying of the parental strand and can pass from one generation to the next via the gametes. The methylating enzyme DNMT1 methylates
preferentially CpG whose partners on the parental strand are already methylated
(see review by Bird 2002).
It is interesting to note that also PRDM14, a PR domain-containing transcriptional regulator is expressed at the PGC specification. Its action is critical in two
events: the reacquisition of pluripotency and successful genome wide epigenetic
reprogramming. In mice Prdm14 mutants, the failure of these events manifests even
in the presence of PRDM1 (BLIMP1), a key transcriptional regulator for PGC
specification (Yamaji et al. 2008). De novo methyltransferases DNMT3A and
DNMT3B are highly expressed during epigenetic reprogramming (see review by
Bird 2002).
Differentially methylated regions of imprinted genes are conserved in migrating
PGCs (Hajkova et al. 2002). In females only one X chromosome is active in specified PGCs. The reactivation of one X occurs during PGC migration and is ­completed
in all PGCs after their arrival in the developing ovary and before their entering into
44
G.H. Vázquez-Nin et al.
meiosis. Reversion of imprinted X chromosome inactivation in the inner cell mass
of the female blastocyst is initiated by the repression of Xist from the paternal X
chromosome. Three of the key factors supporting pluripotency, Nanog, Oct3/4 and
Sox2, bind to Xist intron 1 in undifferentiated embryonic stem cells. The drastic
release of all three factors triggers a rapid ectopic accumulation of Xist RNA and X
inactivation in the epiblast. These three factors cooperate to repress Xist and thus
couple X inactivation reprogramming to control of pluripotency during embryogenesis in mammals (Navarro et al. 2008).
Studies carried out on mice showed that the differentially methylated regions of
maternally or paternally imprinted genes are progressively and synchronously
­demethylated in PGCs. The repetitive DNA elements studied so far are coordinately
demethylated in the male and female mice genomes. Imprinted sequences are
­completely demethylated and repeat sequences only lose some methylation, retaining
a substantial degree of methylation (see review by Lees-Murdock and Walsh 2008).
In mice, the female germline demethylation continues past e15.5 as the female
PGC, now known as primary oocytes, progress through prophase of meiosis (LeesMurdock et al. 2003).
The mechanisms of demethylation have not been clarified. There are at least
two different procedures: active or enzymatic demethylation and slower replication-coupled demethylation. The relationships of demethylase complex to
MBD2b remain unclear. Some works demonstrate that the 5-methylcytosine was
replaced by cytosine during demethylation reaction. Other enzymatic complexes
of hydrolyses 5-methylcytosine to cytosine and methanol (Wolffe and Wade
1999). However, biochemical data suggest this reversal of the methylation is
energetically unfavorable (see review by Lees-Murdock and Walsh 2008).
Replication-coupled demethylation propose that DNA replication can also contribute to eliminating 5-methylcytosine from DNA. If some factor prevents the
access of the methyltrasnferase enzyme to these sites, progressive demethylation
of selected sites can take place (Wolffe and Wade 1999). The mechanism of demethylation in mammals has not been confirmed and no reproducible ­demethylation
activity has been identified (see review by Lees-Murdock and Walsh 2008).
Recent results obtained in mouse support a role of the Activation-Induced cytidine Deaminase (AID), a member of the cytidine deaminase APOBEC family,
and of Base Excision Repair (BER) pathway in genome-wide DNA demethylatioin occurring in PGCs after gonadal ridge colonization (Popp et al. 2010;
Hajkova et al. 2010).
Following erasure of imprints and partial erasure of methylation on repeats,
these sequences must undergo de novo methylation during subsequent germ cell
development to achieve the methylation patterns of the mature gametes (see review
by Lees-Murdock and Walsh 2008). While in the male germ line resetting of
methylation occurs before meiosis, in the female germ line the imprint control
regions are hypomethylated until after the pachytene stage of meiosis I, which
occurs in the postnatal growing oocyte (Lees-Murdock et al. 2005; see also the
review by Lees-Murdock and Walsh 2008).
2 Embryonic Development of the Ovary, Sexual Reproduction and Meiosis
45
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