MEIOTIC PROPHASE PROGRESSION AND
GERM CELL ELIMINATION
IN FETAL AND NEONATAL MOUSE OVARIES
© Adriana Cristina Ene
Department of Biology
McGill University
Montreal, Quebec
Canada
A thesis submitted to McGill University in partial fulfillment of the
requirements of the degree of Master of Science
December 2009
Abstract
In most mammalian species, all oogonia cease mitotic proliferation and enter
meiosis in fetal ovaries. Furthermore, more than half of the maximum number of germ
cells is eliminated from ovaries by neonatal life, thus limiting the oocyte reserve for
reproduction. The cause or mechanism of this female germ cell loss remains largely
unknown. A major loss occurs in the oocytes which reach the pachytene stage of meiotic
prophase, suggesting that oocytes with meiotic or recombination errors may be
eliminated by a checkpoint mechanism. It remains to be determined whether oocytes are
eliminated by apoptosis and if so in which pathway. The purpose of my study is to
investigate a mechanism of oocyte loss in the mouse ovary during meiotic prophase. We
used an Msh5 null mutant mouse strain, in which all oocytes are eliminated by neonatal
life. Msh5 encodes a protein required for meiotic chromosome synapsis.
Msh5 heterozygous mutant mice were crossed and ovaries were isolated from
female progeny at 14.5 – 22.5 days postcoitum (dpc). We studied the loss of germ cells in
Msh5 -/- (MT) females comparing to the Msh5 +/+ (WT) and Msh5 (+/-) (HT) females
by immunolabeling of ovarian sections for GCNA1 or MVH (both germ cell markers) or
by counting GCNA1 positive germ cells in cell suspension preparations. Our results
showed a continuous loss of GCNA1 positive cells in both MT and WT although the loss
in MT was constantly larger than in the WT. A significant difference between WT and
MT was found at 19.5 dpc.
Meiotic progression was studied by GCNA1 and SC (synaptonemal complex) or
SC and ɣH2AX double immunolabeling of chromosome spread preparations. We found
that meiosis in MT was blocked at zygotene-pachytene transition. No normal pachytene
was observed in MT.
The role of apoptosis in elimination of oocytes during meiotic prophase was
investigated by analyzing the cleavage of various caspases (caspase 2, 3, 6, 7, 9) as well
as PARP1 by western blot using the lysate of whole ovaries. The activation of initiator
caspase 9 increased from 17.5 to 18.5 dpc and decreased by 19.5 dpc. Caspase 2L
activation also increased in a similar pattern but at much lower levels. The activation of
effector caspase 3 or 6 remained at low levels. The activation of caspase 7 also was low
ii
but increased slightly at 19.5 dpc. The cleavage of PARP1 was high at all investigated
stages. There were not major differences in the average level of activation between WT
and MT. By immunolabeling of ovarian sections we observed that cleaved caspases and
PARP1 were localized in oocytes but also in cells negative for GCNA1.
These results suggest that a mitochondrial pathway of apoptosis may play a role in
the elimination of oocytes during meiotic prophase, involving activation of caspase 9 and
cleavage of PARP1. However further studies are necessary for identification of an
effector caspase.
iii
Résumé
Dans la plupart des espèces de mammifères, tous les oocytes cessent la
prolifération mitotique et initialisent la méiose dans les ovaires fœtales. En outre, plus de
la moitié du nombre maximal de cellules germinales est éliminée dans les ovaires pendant
la vie néonatale, limitant ainsi la réserve d'oocytes pour la reproduction. La cause ou le
mécanisme de cette perte de cellules germinales femelles reste largement inconnu. Une
perte majeure se produit dans les oocytes qui atteignent le stade pachytène de la prophase
méiotique, suggérant que les oocytes avec des erreurs dans la méiose ou des erreurs de
recombinaison peuvent être éliminés par un mécanisme de contrôle. Il reste à déterminer
si les oocytes sont éliminés par apoptose, et si oui, par quel méchanisme. Le but de mon
projet est d'étudier un mécanisme de perte d'oocytes dans les ovaires de souris durant la
prophase méiotique. Nous avons utilisé une souche de souris mutantes pour la gene
Msh5, dans lequelles tous les oocytes sont éliminés durant la vie néonatale. Msh5 code
pour une protéine nécessaire à la synapse de chromosomes méiotiques.
Des souris hétérozygote Msh5 ont été croisées et les ovaires ont été isolées de la
progéniture féminine de 14,5 à 22,5 dpc. Nous avons étudié la perte de cellules
germinales dans les ovaires des femelles Msh5 -/- (MT) en les comparant à ceux des
femelles Msh5 +/+ (WT) et Msh5 +/- (HT) par immunodétection en utilisant des
anticorps anti-GCNA1 et anti-MVH (marqueurs des cellules germinales) ou par le
comptage des cellules positives au GCNA1 dans des suspensions cellulaires. Nos
résultats montrent une perte continue de cellules positives au GCNA1 chez les souris MT
et WT, bien que la perte chez les MT a été constamment supérieure à celle des WT
(différence significative à 19.5 dpc).
La progression de la méiose a été étudiée par immunodétection double pour
GCNA1 et SC (complexe synaptonémal) ou pour SC et γH2AX sur des préparations de
chromosomes. Nous avons constaté que la méiose chez les souris MT est bloquée dans le
stage de transition zygotène-pachytène. Nous n’avons pas observé de pachytène normal
chez les souris MT.
Le rôle de l'apoptose dans l'élimination d'oocytes au cours de la prophase méiotique a été
étudié par analyse du clivage de diverses caspases (caspases 2, 3, 6, 7, 9) ainsi que celui
iv
de la PARP1 par immunobuvardage des protéines d'ovaires entières lysées. L'activation
de la caspase 9 initiatrice a augmenté entre 17.5dpc et 18.5 dpc et a ensuite baissé à 19.5
dpc. Celle de la caspase 2L a augmenté d'une manière semblable, mais à des niveaux
beaucoup plus bas. L'activation des caspases effectrice 3 et 6 est demeurée à des niveaux
faibles mais celle de la caspase 7 bien que faible a augmenté légèrement à 19.5 dpc. Le
clivage de PARP1 était élevé dans tous les stages. Dans tous ces cas, il n'y a pas eu de
grandes différences dans le niveau moyen d'activation entre WT et MT. Par
immunodétection de sections d'ovaires, nous avons observé que les caspases et PARP1
clivées étaient localisées dans des oocytes, mais aussi dans les cellules sans marquage
pour GCNA1.
Ces résultats indiquent que la voie mitochondriale de l'apoptose peut jouer un rôle
dans l'élimination d'oocytes au cours de la prophase méiotique, puisque les clivages de la
caspase 9 et de PARP1 y sont associés. Cependant des études supplémentaires sont
nécessaires pour l'identification de caspases effectrices.
v
Acknowledgements
I thank Dr. Taketo for offering me the opportunity to work on oocytes. I thank her for
guidance throughout my study, insightful cristicism and financial support. She made my
dream turn into reality. I admire her strength, courage and dedication to science. I
enjoyed our lab discussion!
I thank my spouse, Florian, for his positive attitude, support and encouragement during
my study. Thank you for helping and trusting me!
I am grateful to members of my graduate supervisory committee, Dr. Monique Zetka and
Dr. Hugh Clarke for providing constructive criticism and ideas for further experiments.
I thank Dr. Naumova (Department of Obstretics and Gynecology) for redesigning the
primers for Msh5 genotyping.
I would like to express my gratitude to members of Urology and Surgical Research from
Royal Victoria Hospital for help and support. I am especially grateful to Emmanulle
Devemy for advice and technical help on western blot, for making herself available
whenever I needed help and for being such a nice person.
I am grateful to Eve
Delamirande for advice on western blot experiments, for teaching me with so much
patience how to use the software to quantify my blots and for helping me with translation
in French. Michèle Villemure taught me many procedures in the first part of my project
and she made herself available anytime I needed her advice, thank you Michèle! I thank
Natalya Karp and Krista Zeidan for helping with some of genotyping experiments. Maria
San Gabrielle helped me with various technical advices; I thank her for this and for her
wonderful smile. I also thank Geneviève Lamothe for offering me her help and detailed
explanation whenever I need it.
I thank to all people that helped and supported me during this study.
vi
Table of contents
Abstract..............................................................................................................................II
Résumé..............................................................................................................................IV
Acknowledgments.............................................................................................................VI
Table of contents..............................................................................................................VII
List of figures……………………………………………………………………………..X
List of tables………………………………………………………………………….....XII
List of abbreviations…………………………………………………………………....XIII
1. Introduction…………………………………………………………………...............1
1.1.Oogenesis in mice……………………………………………………………………..1
1.1.1. Origin of the mouse germ cells…………………………………………………….1
1.1.2. Germline-somatic interaction………………………………………………………2
1.2. Meiosis………………………………………………………………………………..4
1.2.1. Meiosis in mammals – cell cycle events in mouse oogenesis…………………….4
1.2.2. Meiotic prophase I……………………………………………………………….....4
1.2.3. Sexual dimorphism in the onset of meiosis I………………………………………7
1.2.4. Molecular mechanism of meiotic prophase I ……………………………………...7
1.2.5. Meiotic checkpoints in mice…………………………………………….................9
1.2.6. Recent opinions on connection between asynapsis and meiotic impairment……..10
1.3. Germ cell loss during ovarian development………………………………………...12
1.3.1. Early observation on germ cell loss and degeneration……………………………12
1.3.2. Causes for germ cell loss……………………………………………….................13
1.4. Apoptosis- caspase dependant and independent pathways…………………….……15
1.4.1. Definition of apoptosis…………………………………………………….….…...15
1.4.2. Caspases – important players during apoptosis……………………………….…..15
1.4.3. Mechanisms of apoptosis………………………………………………………….16
1.4.4. Poly (ADP-ribose) polymerase 1 (Parp 1)……………………………………..….18
1.4.5. Apoptosis of oocytes……………………………………………………………....20
1.4.6. Other mechanisms for oocyte elimination………………………………………...21
1.4.6.1. Caspase independent pathways………………………………………................22
vii
1.5. Experimental design…………………………………………………………………24
1.5.1. Msh5 mutant mouse……………………………………………………………….24
1.5.2. Aim 1. Changes in the number of germ cells in fetal and neonatal ovaries……….25
1.5.3. Aim 2. Analysis of meiotic prophase progression………………………………..26
1.5.4. Aim 3. Study of apoptosis………………………………………………………...26
2. Materials and Methods……………………….…………………………………..….27
2.1. Animals………………………………………………………………………...........27
2.2. Genotyping…………………………………………………………………………..27
2.2.1. Separate amplification of wild-type and mutant alleles………………………….27
2.2.2. Duplex PCR……………………………………………………………………….27
2.3. Isolation of ovaries…………………………………………………………………..28
2.4. Fixation of ovaries for histological preparation…………………………………….28
2.5 Paraffin embedding and microtome sectioning of fixed ovaries…………………..28
2.6. Deparaffinization and hydration………………………………………………….…29
2.7. Antigen retrieval………………………………………………………………….…29
2.8. Dissociated cells and chromosome spread preparations on histological slides by
cytospin centrifugation………………………………………………………………29
2.9. TdT-mediated dUTP nick end labelling (TUNEL)……………………………….....30
2.10. GCNA1and MVH immunolabelling of ovarian sections…………………………..31
2. 11. Double immunolabelling of GCNA1 and ɣH2AX of ovarian sections…………...29
2. 12. Double imunolabelling of GCNA1 and cleaved Caspase 9 (37/15 KDa or 39 KDa)
or cleaved PARP1 (24KDa) of ovarian sections………………………………………....31
2. 13 Immunocytochemical staining of GCNA1/SC, SC/ɣH2AX and GCNA1/SC/ɣH2AX
of chromosome spreads…………………………………………...................................31
2.14. Counting and staging of germ cells………………………………………………..34
2.15 Statistical analysis…………………………………………………………………..34
2. 16. Western Blot………………………………………………………………............34
2. 17. Specificity of antibodies…………………………………………………………..35
2. 17. 1. Blocking peptide………………………………………………………………..35
2. 17. 2. Jurkat Apoptosis Cell Lysates…………………………………………………..35
2. 18. Analysis of band intensity……………………………………………………..….36
viii
3. Results………………………………………………………………………………...37
3.1. Changes in the number of germ cells in fetal and neonatal ovaries……………….37
3.1.1. The abundance of germ cell in ovarian sections detected by GCNA1 and MVH
immunolabelling………………………………………………………………………....37
3.1. 2. Changes in the number of GCNA1 positive germ cells examined in spread cell
preparations………………………………………………………………………............41
3.2. Meiotic prophase progression in WT, HT and MT ovaries………………................42
3.2.1. Identification of meiotic stages using SC/GCNA staining…………………….… 42
3.2.2. Identification of meiotic stages using SC/ ɣH2AX staining……………………...43
3.2.3. Observation on meiotic progression in ovarian sections…………………….……51
3.3. Apoptosis…………………………………………………………………………....53
3.3.1. TUNEL assay……………………………………………………………………...53
3.3.2. Western Blot analysis of the activation of caspase 2, 3, 6, 7, 9 and cleavage of
PARP 1…………………………………………………………………………………..57
3.3.2.1. Caspase 9………………………………………………………………………..57
3.3.2.2 Cleaved Caspase 9 of 35/15 KDa……………………………………………….57
3.3.2.3. Caspase 2 L……………………………………………………………………...57
3.3.2.4. Caspase 3, 6 and 7………………………………………………………........... 58
3.3.3.5. PARP 1…………………………………………………………………………..58
3.3.4. Localization of cleaved Caspase 9 (35/15 KDa and 37 KDa), cleaved Caspase 7 (20
KDa) and cleaved PARP1 (24 KDa) in ovaries at 18.5 dpc………………………….….66
3.3.4.1. Localization of 35/15 KDa cleaved form of Caspase 9…………………............66
3.3.4.2. Localization of 37 KDa of the cleaved form of Caspase 9……………………...66
3.3.4.3. Localization of cleaved Caspase 7……………………………………………....66
3.3.4.4. Localization of cleaved PARP1 (24 KDa)……………………………………....67
4. Discussion…………………………………………………………………………..…72
4.1. Variation in the number of germ cells in fetal and neonatal ovaries during meiotic
prophase progression…………………………………………………….............72
4.2. Apoptotic pathway…………………………………………………………………..73
5. Conclusion……………………………………………………………….…………...75
6. Literature………………………………………………………………………….….76
ix
Figures
Figure 1. Oogenesis and folliculogenesis during early development of the mouse ovary ..3
Figure 2. Meiotic prophase I………………………………………………………………6
Figure 3. The intrinsic and the extrinswic pathways of apoptosis……………………….19
Figure 4. Immunolabelling of ovarian sections for GCNA1 or MVH……………….39-40
Figure 5. Changes in the total number of GCNA-1 positive cells from ovaries…………41
Figure 6. Meiotic progression during ovarian development in WT/HT identified by
GCNA1/SC immunolabelling……………………………………………………47
Figure 7. Meiotic progression during ovarian development in MT identified by
GCNA1/SC immunolabelling……………………………………………………47
Figure 8. Meiotic progression during ovarian development in WT/HT identified by
SC/ɣH2AX immunolabelling…………………………………………………….48
Figure 9. Meiotic progression during ovarian development in MT identified by
SC/ɣH2AX immunolabelling…………………………………………………….48
Figure 10. Meiotic stages in spread chromosomes from WT ovary double immunolabeled
for SC and ɣH2AX ……………………………………………………………...49
Figure 11. Meiotic stages in spread chromosomes from MT ovary double immunolabeled
for SC and ɣH2AX ……………………………………………………………...50
Figure 12. Double immunolabelling of ovarian sections for GCNA1 and ɣH2AX……..52
Figure 13. Distribution of TUNEL positive cells in ovarian sections………………..54-56
Figure 14. Immunoblotting and activation of Caspase 9………………………………...59
Figure 15. Immunoblotting of 35/15 KDa cleaved form of caspase 9…………………...60
Figure 16. Specificity of Caspase 2L antibody…………………………………………..60
Figure 17. Immunoblotting and activation of Caspase 2L……………………………….61
Figure 18. Immunoblotting and activation of Caspase 3………………………………...62
Figure 19. Immunoblotting of and activation Caspase 6………………………………...63
Figure 20. Immunoblotting and activation of Caspase 7……………………………...…64
Figure 21. Immunoblotting and cleavage of PARP……………………………………...65
Figure 22. Double immunolabelling of GCNA1 and the 35/15 KDa cleaved form of
Caspase 9 in ovarian sections……………………………………………………68
x
Figure 23. Double immunolabelling of GCNA1 and the 37 KDa cleaved form of Caspase
9 in ovarian sections………………………………………………………………….69-70
Figure 24. Double immunolabelling of GCNA1 and the 24 KDa cleaved form of Parp in
ovarian sections…………………………………………………………………………..71
xi
Tables
Table 1. Reagents used for immunofluorescent staining…………………..……………32
Table 2. The source of antibodies used for immunofluorescent staining……………….33
Table 3. Specification of antibodies used for western-blot ……………………………...36
Table 4. The number of WT (HT) / MT ovaries investigated using different methods….37
Table 5. Criteria used to define meiotic stages in WT…………………………………...45
Table 6. Criteria used to define meiotic stages in MT…………………………………...46
xii
List of abbreviations
AIF
Apoptosis inducing factor
APAF-1
Apoptotic protease activating factor 1
ATM
Ataxia telangiectasia mutated
ATR
Ataxia telangiectasia and Rad 3 related
BAX
Bcl-2 associated X protein
Bcl-2
B-cell CLL/lymphoma 2; family of genes with pro- or antiapoptotic
role
BRCA1/2
Breast cancer 1/2
CARD
Caspase recruitment domain
CRADD/RAIDD
Caspase and RIP adapter with death domain
D
Diplotene
DED
Death effector domain
DMC1
Disrupted meiotic cDNA
DSBs
Double strand breaks
FADD/Mort1
Fas-Apo 1 associated death domain protein
FAS
Death receptor
FADD
Fas-associated protein with death domain
FasL
Fas Ligand
GCNA1
Germ cell nuclear antigen 1
H2AX
H2A histone family, member X
HOP2
Homologous pairing protein 2
HRR
Homologous recombinational repair
HT
Msh5+/- female mouse
IAP
Inhibitor of apoptosis
L
Leptotene
MEI1
Meiosis defective 1
MLH1
Mutant L homologue 1
MRE11
Meiotic recombination 11
xiii
MSCI
Meiotic sex chromosome inactivation
MSH4
Mutant S homologue 4
MSH5
Mutant S homologue 5
MSUC
Meiotic silencing of unpaired chromatin
MT
Msh5-/- mouse female
MVH
Mouse vasa homologue
NBS1
Nibrin protein (associated with DSBs repair)
OMI/HtrA2
Serine threonine HtrA2, mitochondrial protein (High Temperature
requirement protein or Omi stress-regulated endoprotease)
P
Pachytene
P53
Protein 53
PARP1
Poly(ADP-Ribose) Polymerase-1
PCD
Programmed cell death
PGCs
Primordial germ cells
PMS2
Post-meiotic segregation increase 2 protein
RAD50
DNA repair protein RAD50
RAD51
Rec-A like protein
SC
Synaptomal complex
Smac/Diablo
Second mitochondria-derived activator of caspases
SPO11
Meiosis specific protein SPO 11 (Sporulation-specific protein 11)
SYCP1
Synaptomal complex protein 1
SYCP2
Synaptomal complex protein 2
SYCP3
Synaptomal complex protein 3
TUNEL
TdT-mediated dUTP nick end labelling
Z
Zygotene
WT
Msh5+/+ mouse female
xiv
Introduction
1.1 Oogenesis in mice
1.1.1. Origin of the mouse germ cells
Germ cells are defined as those cells, all of whose surviving descendants will
become sperm or eggs (McLaren 2003). In the mouse, primordial germ cells (PGCs)
form far from the site of the developing gonads and migrate into the developing gonads.
PGCs can first be detected at 7.5 dpc as a cluster of alkaline phosphatase-positive cells at
the base of allantois (Ginsburg et al 1990) when they become lineage restricted. By 9.0
dpc, the posterior visceral endoderm is invaginating to form the hind gut, and the PGCs
from the cluster are carried along with the endoderm and distributed along the ventral
length of the hind gut (McLaren 2000). Between 9-9.5 dpc, PGC move from the ventral
to the dorsal side of the gut and migrate laterally to colonize the genital ridges between
10-11 dpc. It was suggested (Donovan et al. 1986) that once the germ cells arrive at the
gonadal ridge, (10.5-12.5 dpc) they undergo major changes in phenotype: the diameter of
germ cells increases, they become more rounded and less motile both in vivo and in vitro
(Donovan et al 1986) and changes in the appearance of the cytoplasm were observed
(Spiegelman and Bennet 1973). Expression of stage-specific antigen 1 (SSEA-1), first
evident at 9.5 dpc, is downregulated by 12.5 dpc and new proteins appear, including
germ cell nuclear antigen 1 (GCNA-1) (Enders and May 1994) and mouse vasa
homologue (Fujiwara et al. 1994). Once female mouse PGCs enter the genital ridge, they
divide by mitosis until approximately 13.5-14.5 dpc, when about 12.500 germ cells are
present (Tam and Snow, 1981). In this stage PGCs are called oogonia. Cyst growth is
accompanied by incomplete cytokinesis and the dividing germ cells are held together by
intercellular bridges (Gondos 1973). Subsequent sexual differentiation of germ cells is
determined by the gonadal environment that harbors them (Peters 1970, McLaren 1995).
When mitotic proliferation stops in females, oogonia immediately start prophase of
meiosis and become oocytes (Wassarman 1988). Meiosis is arrested at the end of
prophase I soon after birth. Meiotic division is resumed after puberty upon ovulation and
fertilization when two consecutive meiotic divisions take place.
1
1.1.2. Germline-somatic interaction
In female mice, while oogonia are dividing to form cysts, they also interact with
somatic cells in the ovary. The germ cells and epithelial pregranulosa cells become
organized into ovigerous or ovarian cords, which remain until primordial follicles begin
to form (Byskov, 1986, Guigon and Magre 2006) (see Fig. 1). The somatic component of
the ovary appears to have little influence on the onset of meiosis in germ cells. Entry into
meiosis occurs at the same chronological time whether the germ cells are in the ovary, or
ectopically in the developing adrenal, or explanted into a cultured aggregate of
embryonic lung tissue (McLaren and Southee 1997). Around the time of birth, the cysts
break and the somatic cells in the ovary invade clusters of germ cells (Byskov 1986) so
that each surviving oocyte becomes surrounded by a single layer of flattened granulosa
cells, forming the primordial follicle (Fig. 1). The oocytes are arrested in the diplotene
stage of first meiotic prophase, until oocyte growth is initiated due to an unknown
stimulus.
The germ cells have an influence on the somatic cell lineages in the fetal ovary
(McLaren 1991). If only few cells enter the genital ridge (the precursor of the
mammalian gonad), the supporting cell precursors differentiate only to the stage of prefollicle cells which are subsequently lost. The resulted ovary is underdeveloped and
contains only stromal tissue (McLaren 1991). If germ cells enter, the granulosa cell
differentiation is initiated and after the entry in meiosis they will form primordial
follicles around the time of birth. However, if the oocytes are lost, such in the case of
ovarian grafts in adult male mice, the pregranulosa cells are capable of transformation
into Sertoli cells (Taketo-Hosotani et al. 1985, Taketo-Hosotani and Merchant-Larios
1986).
2
Primordial
Germ Cells
Primordial
Follicles
Germline Cyst
Cysts
breakdown
11
13
15
17
19
mitosis
(oogonia)
arrival at
gonad
Germ cells
enter
meiosis
(oocytes)
20
24 dpc
birth
Oocytes enter
meiotic arrest at
diplotene (diploid
oocyte 2n 4C)
Figure 1. Oogenesis and folliculogenesis during early development of the mouse ovary
(adaptated after Pepling 2006)
3
1.2. Meiosis
1.2.1. Meiosis in mammals – cell cycle events in mouse oogenesis
Meiosis is a specialized cell division that is essential for the reproduction of all
sexual organisms. During meiosis a diploid cell (2N) undergoes two rounds of division
after a single DNA replication event to produce haploid gametes for fertilization. During
the first division (meiosis I), also called “reductional division” homologous
chromosomes (paternal / maternal) are separated into two daughter cells. The resulting
cells are haploid (N) and each contains one set of bichromatidic chromosomes (2C). The
second division (meiosis II), also called “equational division”, proceeds without another
round of DNA replication and the sister chromatids of each chromosome segregate into
two daughter cells. Therefore the resulting cells are haploid (N) and each contains one
set of monochromatidic chromosomes (C).
In females, only one haploid cell forms as a result of meiotic divisions and
retains most of the cytoplasm. The other two cells, called “polar bodies”, which result
from the first and the second meiotic division, will degenerate.
Meiosis I is subdivided into four stages: prophase I, metaphase I, anaphase I and
telophase I. Prophase I is a complex process characteristic for meiosis and will be
described in detail in section 1.2.2. During metaphase I the homologues chromosomes
align along an equatorial plate and in anaphase I each chromosome of a pair migrates to
the opposite pole of the cell. In telophase I the two sets of chromosome separate to the
cell poles and two haploid cells are formed.
Meiosis II is similar to mitosis. Chromosomes align along an equatorial plate
during meiosis II and in anaphase II each sister chromatid migrates to the opposite poles
of the cell. In telophase II the cell wall cleaves and two new cells are formed.
1.2.2. Meiotic prophase I
Prophase I is an important feature of meiosis when events such as chromosome
pairing, synapsis and crossing-over take place. Prophase I is divided into five stages:
4
leptonema, zygonema, pachynema, diplonema and diakinesis (reviewed in Cohen et al.
2006, Morelli and Cohen 2005). The different stages of prophase I are defined by several
criteria such as the degree of chromosome condensation and the formation of the
synaptonemal complex (SC). The synaptonemal complex (SC) is a zipper-like protein
structure which consists of two lateral/axial elements which forms along each
homologue and a transversal central element that holds together sister chromatids during
synapsis (Cohen et al. 2006). SC consists of three main proteins, called SC protein
(SYCP or SCP): SYCP1, SYCP2 and SYCP3. SYCP3 is the main constituent of the
lateral/axial element while the central element consists of SYCP1. The formation of SC
proteins can be tracked by immunostaining to identify meiotic stages (Fig. 2).
At leptonema (adjective: leptotene) chromosomes begin to condense and initiate
homologue searching and pairing. The lateral element starts to form along each sister
chromatid (Fig. 2). Another event is the initiation of meiotic DNA double-strand breaks
(DSBs) (Mahadevaiah et al. 2001). Programmed DNA cuts are necessary for the
initiation of homologous recombination. During zygonema (adjectiv: zygotene)
homologous chromosomes align, and synapsis begins with the aid of the central element
(SYCP1) which starts to accumulate (Fig. 2). Progression in synapsis is accompanied by
DSBs repair. The lateral element is fully formed by the end of this stage as a continuous
element and telomeres are tightly clustered in the so-called “bouquet configuration”.
When cells enter pachynema (adjectiv: pachytene), the longest stage of prophase I, the
homologues are completely synapsed. The lateral elements, now called the “axial
elements” are fully joined with the aid of the transverse filaments formed by proteins of
the central element (Fig. 2). This is the stage when crossing-over and genetic
recombination between chromosomes of different parental origin take place thus
ensuring an increase in genetic variability. In diplonema (adjectiv: diplotene) the
homologs start to repel each other but they remain connected at the chiasmata which is
the physical manifestation of crossover (Fig. 2). In this stage the SC begins to dissolve
and by diakinesis, most of the SC structure is lost.
5
LEPTONEMA
Sister chromatids
Centromer
e
ZYGONEMA
Central element
(SYCP1)
Lateral element
(SYCP2, SYCP3)
PACHYNEMA
Axial element
(SYCP2, SYCP3)
DIPLONEMA
Chiasmata
Figure 2. Meiotic prophase I (adapted from Morelli and Cohen 2005)
6
1.2.3. Sexual dimorphism in the onset of meiosis I
The onset as well as continuity of meiosis differs between the two sexes. In female
mice all germ cells initiate meiosis during embryonic development at around 13.5 dpc.
By birth oocytes arrest meiosis at “dictyate” until after sexual maturation when some
oocytes are recruited at each estrous cycle. The selected oocytes go throughout first
division and are arrested in metaphase II. If fecundation takes place, oocytes resume
meiosis and undergo the second division. On the contrary, in males, meiosis is a
postnatal event which initiates before puberty and continues in waves throughout adult
life without an arrest period (reviewed in Morelli and Cohen 2005). Another difference is
the SC length which in females is twice as long as in males. Also, different phenotypes
were observed for male and female mutant mice for genes involved in meiotic
progression.
1.2.4. Molecular mechanism of meiotic prophase I
Recent studies revealed many genes with essential roles in meiosis in mammals
(reviewed by Morelli and Cohen 2005, Cohen et al. 2006). It was observed that different
mutants for genes with roles in SC formation (Sycp3), DSBs formation and repair
(Spo11, Dmc1, Rad 51, Msh4, Msh5), recombination (Mlh1, Mlh3) cause synaptic and
recombination failure in both sexes but there is a difference in the stringency of
phenotypes between males and females. In males, in most of the cases, meiosis cannot
progress beyond late zygotene or pachytene stage which would cause infertility due to
absence of mature spermatozoa (reviewed in Barchi et al. 2005). In contrast, the females
display a broad range of phenotypes from reduced fertility to complete infertility
depending on the nature of defect (Morelli and Cohen 2005).
Homologous recombination is initiated in almost all meiotic species by the
formation of DSBs in DNA strands during the leptotene (Madevaiah et al. 2001). DSBs
are formed by the topoisomerase-like enzyme SPO11. Spo11-/- female mice appear to
have a normal progression of meiotic prophase I but most of the oocytes are lost at or
before primordial follicle formation (Di Giacomo et al. 2005). On the contrary, the
Spo11-/- males arrest at or before pachytene (Baudat et al. 2000, Barchi et al. 2005).
7
After the removal of SPO11 from chromosomes, the DSBs are repaired by
strand invasion and recombination. In eukaryotes this step is catalyzed by RAD51 and
DMC1, two eukaryotic homologues of the bacterial RecA protein. RAD51 forms nuclear
foci from early zygonema colocalizing with the lateral element SYCP3 at distinct loci
called meiotic nodules. DMC1 is restricted to meiosis and colocalizes with RAD51.
Male and female mice homozygous for a null mutation in Dmc1 -/- are sterile and the
meiosis progress apparently normal until zygonema but no normal pachytene is observed
(Pittman et al. 1998).
As synapsis initiates and DSBs are processed, RAD51 and DMC1 are lost from
meiotic nodules and replaced with downstream players in meiotic recombination such as
BRCA1. BRCA1 colocalizes with ATM and ɣH2AX at unsynapsed chromosomes
through zygonema and disappears after synapsis. ATM, the protein product of the gene
mutated in the human autosomal recessive disorder ataxia telangiectasia, and the related
protein, ATR, belong to the PI3-kinase (phosphoinositide-3-kinase) family of proteins
that recognize DNA damage. Both ATM and ATR localize to SC. ATR associates with
unsynapsed lateral elements (Keegan et al. 1996) from leptonema to pachynema and is
mostly gone by mid pachynema. ATM appears along asynapsed and synapsed meiotic
chromosomes from late zygonema to pachynema (Plug et al. 1997). ATM activates
many of the proteins involved in DSB repair including RAD50/MRE11/NBS1 complex
and the BRCA1/2 checkpoint proteins (reviewed by Lavin et al. 2005, Bannister and
Schimenti 2004).
Other proteins involved in meiotic progression are the members of MMR
machinery such as eukaryotic homologs of bacterial MutS (MSH4 and MSH5) and MutL
(MLH1, MLH2, MLH3 and PMS2) (reviewed in Cohen et al. 2006). MSH4 forms
heterodimers with MSH5 and they appear to function only in meiotic recombination.
MSH4 and probably MSH5 start to form in leptonema at the sites of DSBs (Kneitz et al.
2000; Cohen et al. 2006) and colocalize in the first part of prophase with RAD51 and
DMC1 (Neyton et al. 2004). In Msh4-/- and Msh5-/- mice meiosis is disrupted early in
prophase resulting in sterility of both males and females (Edelmann et al.. 1999, de Vries
et al. 1999). The axial elements SYCP3 are formed but the synapsis is severely reduced.
8
MSH4 persists until late pachynema and from early pahynema onward interacts
with the late meiotic nodules components such as MLH1 and MLH3. The heterodimer
MLH1/MLH3 forms on meiotic chromosomes after synapsis occurs and marks the sites
of crossing–over events. Spermatocytes from Mlh1 and Mlh3 null mutants progress
through pachynema normally with complete synapsis being formed (Edelmann et al.
1996, Lipkin et al. 2002). However at diplonema the chromosomes desynapse
prematurely and the chiasmata formation fails. In females the oocyte reaches diplonema
but the lack of chiasmata become evident after meiotic resumption upon ovulation. Most
oocytes fail to achieve the anaphase due to the spindle poles destabilization (Woods et
al. 1999).
These observations suggest that there are mechanisms monitoring the cell cycle
events during meiosis, the so-called “checkpoints” which are less stringent in females
comparing to males.
1.2.5. Meiotic checkpoints in mice
A checkpoint mechanism is defined as a mechanism ensuring the proper order
of events during cell division by arresting or delaying the cycle in response to defects in
cellular processes (Hartwell and Weinert 1989). Hochwagen and Amon (2006) defined
the term checkpoint as a mechanism that allows the coupling of two events and which
comprise several components such as a signal, detected by signal sensors; which in turn
activate signal-transduction pathways that translate the signal into an output.
The existence of checkpoint mechanism has been firstly reported in mitosis, the
so called “G2/M checkpoint” (Hartwell and Weinert 1989). This checkpoint ensures that
cell division does not proceed further if unrepaired DSBs are still present. Checkpoints
also operate in meiosis to avoid deleterious outcomes due to errors in recombination. The
meiotic checkpoint mechanisms are better studied in budding yeast. The meiotic
checkpoint in mice and generally in mammals is not well defined in literature because
the signaling pathway is still unknown. In mammals most of the studies focused on
males. The female meiotic checkpoint was less investigated due to the fact that most of
the meiotic prophase takes place during embryonic life hence is less accessible for study.
Another difficulty in studies of the meiotic checkpoint in mammals is the sexual
9
dimorphism in the stringency of mutant phenotypes. Several meiotic checkpoint
mechanisms have been proposed in yeast (reviewed in Roeder and Bailis 2000,
Hochwagen and Amon 2006) and some of these checkpoints are active also in mice.
One of them is the recombination checkpoint or pachytene checkpoint. Mice lacking
Dmc1, Hop2, or Msh5 (with role in recombination) experience a block in gametogenesis
(De Rooji and de Boer 2003). Inactivation of Spo11 or Mei1 (involved in DSBs
formation) in Dmc1-/- or Msh5 -/- mouse female results in a by-pass of the cell-cycle
arrest at pachytene (Di Giacomo et al 2005, Barchi et al 2005, Reinholdt and Schimenti
2005). This suggests that a checkpoint in mouse detects unprotected DSBs and/or the
persistence of subsequent repair intermediates. However, no components of the mouse
recombination checkpoint have been identified.
A “synaptic checkpoint” has also been proposed to be active in mice.
(Hochwagen and Amon 2006). Mice lacking Spo11 and Mei1 block meiosis in the
absence of DSBs but at a latter moment than the block of Dmc1 or Msh5. Double
mutants Spo11-/-Dmc1-/- and Spo11-/- Msh5-/- show similar phenotypes to Spo11-/mice. This implies that a lack of synapsis is sensed by a different checkpoint pathway
which is independent by DSBs-derived unrepaired recombination intermediates (Di
Giacomo et al. 2005, Barchi et al. 2005, Reinholdt and Schimenti 2005).
1.2.6. Recent opinions on the connection between asynapsis and meiotic
impairment
It has been assumed that the sexual dimorphism in the mutant phenotypes for
genes involved in meiotic progression may be due to less efficient checkpoint/
checkpoints in females. A recent review sugests the existence of a DSBs related
pachytene checkpoint in spermatocytes and proposes a new interpretation for the
different response between males and females (Burgoyne et al. 2009). Efficient synapsis
is dependent on the formation and processing of DSBs. It has been recently established
that in human and mouse the asynapsed chromosomes or chromosome regions are
transcriptionally silenced (Baarendes et al. 2005, Turner et al. 2004, Turner et al. 2006,
Ferguson et al. 2008). This phenomenon, referred to as meiotic silencing of unpaired
chromatin or MSUC, is associated with the accumulation of BRCA1 and ATR on
10
asynapsed axes and the phosphorylation of H2AX. The meiotic sex chromosome
inactivation or MSCI in males may be seen as a particular case of MSUC. This new
hypothesis (Burgoyne et al. 2009) assumes that there is a limited stock of molecules such
as ATR, BRCA, H2AX able to recognize and repair asynapsis. Females with extensive
asynapis will fail to achieve MSUC because ATR and/or BRCA1 are blocked at the sites
of unrepaired DNA and thus will avoid oocyte loss. However, if the DSBs are not
repaired, such in Dmc1 -/- females, the oocyte are lost probably due to a G2/M –related
checkpoint response. In males the consequence of asynapsis is also abrogation of MSUC
response and MSCI failure. The activation of the X chromosome at the pachytene stage
leads to the death of male germ cells. As previously proposed (Burgoyne et al. 2007) the
mid pachytene spermatocytes apoptosis in HRR mutants may be not a manifestation of
an ATM/ATR mediated checkpoint response but rather is a consequence of inappropriate
transcription from the X and Y chromosomes due to MSCI failure.
Another recent paper (Kouznetsova et al. 2009) showed that the MSUC has a
limited function in the quality control mechanism in oocytes. MSUC response occurs in
mouse oocytes with a limited degree of asynapsis (Kouznetsova et al. 2009). MSUC is
abrogated in case of extensive asynapsis, probably to a limited axis-associated BRCA1
pool in oocytes. However, oocytes with an extensive degree of asynapsis are eliminated
after birth suggesting the existence of additional mechanisms of control such as DNA
damage or recombinational checkpoints (Di Giacomo et al. 2005).
11
1.3. Germ cell loss during ovarian development
The oocyte pool is limited by the number of germ cells that enter meiosis and
also by a major loss of germ cells in fetal and neonatal life (McClellan et al. 2003). In
some mammalian species (such as human, rat and mouse), up to two thirds of the
oocytes die early in development before follicle formation (Beaumont and Mandl 1962,
Burgoyne and Baker 1985, McClellan et al. 2003). For example, in human females the
number of oocytes reaches a maximum of 7 million at 5 months of gestation and
decreases to 2 million by birth (Baker 1963). The causes or the mechanisms of female
germ cell elimination remain largely unknown.
1.3.1. Early observations on germ cell loss and degeneration
Early studies focused on identifying cell degeneration during prenatal and
neonatal ovarian development by morphological observations. For example, in rat, four
major waves of degeneration of female germ cells have been described (Beaumont and
Mandl 1962, Franchi and Mandl 1962). The first two waves occur in oogonia before the
onset of meiosis and are morphologically characterized by pyknosis and “atretic
division”. Germ cells belonging to the last category lack a mitotic spindle and have a
slight eosinophilia of the cytoplasm.
The third wave is seen in the germ cells in meiotic prophase particularly at the
pachytene stage. These cells, called “Z cells” are characterized by condensed
chromosomes and eosinophilic cytoplasm. Later, several authors observed “Z cells” in
other species. For example, in humans, one-third of pachytene oocytes have been
described as degenerated (Z-cells) or with synaptic errors (Speed 1988). “Degenerating
pachytene” has been also observed by Borum (1961) in the mouse.
The fourth wave of degeneration in rat has been observed in germ cells with a
chromatin configuration typical of the diplotene stage and with a marked crinkling of the
nuclear membrane. The cytoplasm of these cells is less eosinophilic and condensation of
chromatin is less pronounced comparing to the “Z cells”. The loss of oocytes in rat
starting with the onset of meiosis (17.5 dpc) closely parallels the estimated number of
degenerating germ cells observed with a delay of 24 hours (Beaumont and Mandl 1962).
The authors concluded that the time taken between the onset of degeneration and
12
“disappearance” is of the order of 30 hours. These early observation suggest that the
degeneration and the loss of oocytes occur in waves and it was postulated that genetic
errors (during mitotic proliferation and meiosis before follicle formation) or
environmental factors may be responsible for this loss.
1.3.2. Causes for germ cell loss
In an attempt to identify the cause(s) for oocyte loss subsequent studies used
different experimental approaches such as in vitro cultures of germ cells/ovary or study
of different knockout mouse models.
Based on accumulating bodies of evidence, several hypotheses have been
subsequently forwarded to account for the major germ cell loss (reviewed in Tilly 2001).
The first one is so-called “death by neglect”, according to which germ cells may die due
to a limited amount of trophic factors. It was speculated that germ cell loss may be a
random process that continues into adulthood, exacerbated by nutritional deficits or
environmental factors (Pepling 2006). This hypothesis is partly based on the observation
that the addition of growth factors such as leukemia inhibitory factor (LIF), retinoic acid
(RA) or stem cell factor (SCF) to fetal mouse ovaries in culture prevents germ cell
apoptosis (Morita and Tilly 1999, Pesce et al. 1993). However, there is no evidence that
this is a normal component of oogenesis in vivo (Tilly 2001).
A second hypothesis is “death by defect”. Oocytes with meiotic pairing or
recombination anomalies may be eliminated by a checkpoint mechanism. Speed (1988)
described three major categories of synaptic defect among human oocytes from
chromosomally normal fetuses: partial or complete asynapsis of lateral elements, nonhomologous pairing, duplication or insertion. In the female mouse, the existence of a
checkpoint detecting unrepaired DSBs and/or the persistence of repair intermediates has
been proposed (Di Giacomo et al. 2005). Another line of evidence comes from studies in
female mice which lack an X-chromosome or with an inversion in one X chromosome
(Burgoyne and Baker 1985) showing oogenesis failure probably due to a defect in
chromosome pairing. However, it is still uncertain to what extent the genetic defects
contribute to oocyte loss in normal development (De Felici et al 2008).
13
A third theory is “death by self-sacrifice” or the “altruistic death” (Tilly
2001). A major germ cell loss was reported in neonatal mice when cysts start to
breakdown (Pepling and Spradling 2001). According to this theory the “nurse” oocytes
within a cyst may die in order to donate their organelles and cytoplasm to the surviving
oocyte. However, this theory is based mainly on studies on Drosophila and Xenopus
laevis. Another hypothesis based on observations in mice proposes that germ cell
apoptosis within a cyst may be related with an evolutionary conserved process of
mitochondrial selection (Pepling and Spradling 2001). According to this hypothesis,
mitochondria with functional and defective genomes would be actively transported into
different germ cells and the quality of each cell would determine if it survives or enters
apoptosis (Pepling and Spradling 2001). However, in the last case germ cell loss is due
to a genetic defect and cannot be regarded as “death by self-sacrifice”.
A recent study on mouse oocytes throughout meiotic prophase showed a
continuous loss of oocytes during fetal development without evident peaks of
degeneration, suggesting that multiple causes are responsible for oocyte loss at different
stages of development (McClellan et al. 2003).
Most of the conclusions/hypotheses regarding the causes of germ cell loss are
still based on circumstantial evidences. What triggers the loss of oocytes is still unclear,
making it difficult to conclude if oocyte elimination is a random/stochastic process or if
it is governed by evolutionary mechanisms of selection within a developmentally
regulated process. These difficulties are due, at least in part to the fact that ovarian
development is accompanied by complex biologically processes such as mitotic
proliferation, meiosis, follicle formation etc. which are not synchronized events and
overlap to a certain degree. The possibility that a delay may occur between the beginning
of “degeneration” and the actual loss in germ cell number makes any interpretation even
more difficult. Another major drawback is that the mechanism/cell signaling pathway is
not elucidated, making it difficult to establish a cause effect relation.
14
1.4. Apoptosis- caspase dependent and independent pathways
1.4.1. Definition of apoptosis
Apoptosis or programmed cell death (PCD) is an important mechanism of
maintaining homeostasis during development and for responses to external stimuli in
multicellular organisms (Jacobson et al. 1997). The initial description of apoptosis was
based on morphological features including cytoplasm shrinkage, membrane blebbing,
chromatin condensation into dense masses located next to the nuclear envelope and
nucleus and cytoplasm fragmentation. The result is the formation of so-called “apoptotic
bodies” which are degraded by lysosomes in the adjacent cells. Unlike necrosis, there is
no inflammation and the apoptotic cells are eliminated without disruption of the
surrounding tissue. Several biochemical and immunohistochemical detection methods of
apoptosis were subsequently introduced. Wyllie (1980) described fragmentation of
nuclear DNA into multiples of 180 bp as the result of endonuclease activation. When
fragmented DNAs were electrophoresed in an agarose gel, they separated into a
characteristic DNA ladder pattern.
1.4.2. Caspases – important players during apoptosis
In mammals, an important role in apoptosis is played by a family of intracellular
cysteine protease known as caspases or Cysteine Aspartyl-specific Proteases (Reed
2000). They also play a role in inflammation (reviewed by Cohen 1997, Los et al.1999,
Nicholson and Thornberry 1997). Caspases act in a proteolytic cascade whereby they can
activate themselves and each other and they can be classified as upstream (initiator)
caspases or downstream (effector or executioner) caspases (Salvesen and Dixit 1997,
Reed 2000). The initiators are responsible for processing and activation of the
executioners. The executioners are responsible for proteolytic cleavage of a number of
cellular proteins leading to the characteristic morphological changes and DNA
fragmentation (Cohen 1997, Porter et al. 1997, Salvesen and Dixit 1997).
Caspases are present as inactive zymogens in all animal cells and contain a
variable length prodomain and two catalytic subunits known as the large and the small
subunits. The proforms of upstream initiator caspases possess large N-terminal
15
prodomains which function as adaptor proteins allowing interaction with various
molecules that trigger caspase activation (Reed 2000). Large prodomains contain protein
motifs such as CARD (caspase recruitment domains, found in Caspase-9 and -2) or DED
(dead effector domains, found in caspase 8 and 10) that are homologous to motifs
present in a number of signaling molecules such as FADD/Mort 1, CRADD/RAIDD and
Apaf-1 (Boldin et al. 1996, Ahmad et al. 1997, Duan and Dixit 1997). The proforms of
downstream effector caspases contain only short N-terminal prodomains with no
apparent function. Caspases can be activated by cleavage at aspartic acid (Asp) residues
generating a large (approx. 20 KDa) and small (approx 10 KDa) subunit of the active
enzyme.
14 caspases have been identified in humans and mice. Experiments on caspase –
deficient mice (caspase 1, 2, 3, 7, 8, 9 and 11 knockouts) showed cell death
abnormalities before and after birth (reviewed by Zheng et al. 1999; Johnson and
Bridgham 2002). These knockouts exhibit tissue- and cell type-specific or stimulusdependent defects of apoptosis suggesting that different sets of caspases are involved in
separate cell death pathways in vivo. It was shown for example that Caspase 9 knockout
mice die perinatally with enlarged and malformed cerebrum caused by reduced apoptosis
during brain development (Kuida et al. 1998). Experiments using caspase 2 and 9 double
knockout cells demonstrated that these cells can undergo apoptosis triggered by various
stimuli which suggests that other caspases might act redundantly (Zhihotovski and
Orrenius 2005). These studies also suggested that caspases might play an important role
in oocyte apoptosis/elimination.
1.4.3. Mechanisms of apoptosis
The mechanism of apoptosis is evolutionarily conserved. Three major apoptotic
pathways have been described in mammals. The first one, called the intrinsic or
mitochondrial pathway, is triggered by extracellular or intracellular factors such as DNA
damage (Fig. 3). This pathway involves the participation of mitochondria, which release
proteins such as cytochrome c into the cytosol. Cytochrome c binds to Apaf, in the
presence of ATP, which will in turn binds procaspase 9, creating the apoptosome which
is able to cleave the initiator caspase 9. It was shown (Srinivasula et al. 1998) that
16
processing of procaspase-9 occurs by an intrinsic autocatalytic activity of procaspase-9
itself, and that Apaf-1 triggers this activity by oligomerizing procaspase-9. Processing at
Asp-315 activates caspase 9 producing a 35KDa fragment.
The second pathway (Fig. 3), also known as the the extrinsic or death receptor
pathway, involves ligation of the death receptor (such as Fas) to its ligand, FasL. Binding
of FasL to Fas induces trimerization of Fas receptors, which recruit FADD (Fasassociated death domain) through shared death domains (DD). FADD also contains a
“death effector domain” or DED in its N-terminal region. The FAS/FADD complex then
binds to the initiator caspase 8 or 10 (Hikim et al. 2003). A third pathway involves the
endoplasmic reticulum (ER) and initiator caspase 12. This pathway is stimulated by the
accumulation of unfolded or misfolded proteins which result in inhibition of translation
of induction of gene expression (Orrenius et al. 2003).
Crosstalk between these pathways occurs at some levels. Activation of the
upstream caspases converge on the effector caspases such as caspase 3, 6, 7. For
example, after activation, caspase-9 can initiate a caspase cascade by directly activating
procaspase-3 and 7 and indirectly activating procaspase-6. Activated caspase 3 can feed
back on procaspase 9 and activate it by processing its precursor at Asp-330, generating a
37 KDa fragment (p37a) (Srinivasula et al. 1998). This process may have a role in
amplifying the caspase cascade. Finally, the activation of effector caspases will produce
alterations within the nucleus by cleaving a set of nuclear proteins (such as PARP1,
lamin, actin etc) and DNA fragmentation.
Caspase activation and activity are regulated by pro- and anti-apoptotic proteins
such as members of the Bcl2-related family and IAPs (inhibitor of apoptosis proteins).
The Bcl-2 family of proteins plays an important role in the mitochondrial pathway, with
proteins such as Bax functioning as promoters and proteins such Bcl-2 as suppressors of
cell death (Adams et al. 1998, Reed 2000). IAPs (inhibitor of apoptosis) represent a
family of evolutionarily conserved apoptosis suppressors (Deveraux and Reed 1999).
They have an important role in controlling the catalytic function of cytochrome c
(Bröker et al. 2005). Proteins such XIAP, cIAP1 and cIAP2 are able to inhibit initiator
caspase-9 and effector caspases 3 and 7 (Deveraux et al. 1997, Roy et al. 1997,
Deveraux et al. 1999). The IAP proteins are controlled by two other mitochondrial
17
proteins, Smac/Diablo and OMI/HtrA2 (Bröker et al. 2005) which are released after proapoptotic stimuli.
1.4.4. Poly (ADP-ribose) polymerase 1 (PARP1)
The Parp1 gene encodes a ubiquitous enzyme thought to have a critical role in
genome stability and in repair of DNA damage caused by genotoxic exposure
(Vidakovic et al. 2005). Parp1 is activated in response to DNA single strand breaks to
synthesize protein-bound ADP-ribose polymers from NAD+. The enzymatic DSBs
response network described in mammalian germinal cells involves RAD51, ATM, DNAPK and H2AX which are regulated by tumor suppressors like p53 (Richardson et al
2004). Excessive DNA damage generates large branched-chained PAR polymers, which
leads to the activation of a cell-death program (Yu et al. 2002). Cleavage of PARP1 is
one of the late events during apoptosis.
18
Intrinsec pathway
Extrinsec pathway
Ligand
DNA damage
Caspase 2 ?
Death Receptor
Mitochondria
Apoptosome
Adapter molecule
Caspase 9
Caspase 8
Cleaved
caspase 9
(35 KDa)
Cleaved
caspase 9
(37 KDa)
Caspase 3, 6, 7
PARP
Caspase 2 ?
DNA fragmentation
Cell death
Figure 3. The intrinsic and the extrinsic pathways of apoptosis
19
1.4.5. Apoptosis of oocytes
The process of oocyte apoptosis is still poorly documented comparing to follicle
apoptosis (Reynaud and Driancourt 2000). Several studies indicate that PCD in fetal
mouse and human oocytes occurs by apoptosis (reviewed in Reynaud and Driancourt
2000, De Felici 2005). Coucouvanis et al. (1993) identified apoptotic cells in fetal mouse
ovaries by flow cytometry. Apoptotic cells were identified at 13 dpc (4.3%) but an
increased frequency of apoptotic cells was observed at 15 and 17 dpc (8.7-11.3%). The
sorted cells were positive for alkaline phosphatase staining confirming that they are germ
cells. Ratts et al. 1995 reported the presence of DNA ladders at 15.5, 18.5 and birth. At
13.5 dpc the DNA ladders were absent but it was not possible to distinguish if apoptosis
occurred in germ cells or somatic cells. In 1997 De Pol et al. have shown, using the
TUNEL assay, that apoptosis in 18-week-old human fetuses is restricted to germ cells
(8%). It was suggested that a loss of female germ cells may occur throughout the meiotic
prophase at the zygotene/pachytene (13 – 16 dpc) and also at the time of primordial
follicle formation (18 dpc to birth).
Cultured oocytes undergo oligonucleosomal DNA and PARP1 cleavage, two
markers for late apoptosis (Lobascio et al. 2007). Another study performed in vivo
showed that oocytes in meiotic prophase I during fetal and neonatal life are positive for
TUNEL and cleaved PARP1 (Ghafari et al. 2007).
It was suggested that caspase 2 may play a role in both positive and negative
regulation of cell death (Zhivotosky and Orrenius 2005) via two isoforms, caspase 2L
and caspase 2S derived by alternative splicing (Wang et al. 1994, Kumar et al. 1997).
Caspase 2L induces cell death while caspase 2S, a truncated version of caspase 2L
lacking the p19 region, can antagonize cell death. Caspase 2L is the only isoform found
in the ovary (Bergeron et al. 1998). It has been proposed that caspase 2 may have an
effector role in the mitochondrial pathway, being the preferred substrate for caspase 3
(Paroni et al. 2001). On the other hand, Robertson et al. 2002 proposed that nuclear
caspase 2 is an important upstream promoter of cytochrome c release in DNA damageinduced apoptosis via the mitochondrial pathway. They showed that the proteolytic
caspase cascade in response to a low dose of etoposide is as follows: caspase 2
20
mitochondria  caspase 9  caspase 3. Unlike initiator caspases 8 and 9, caspase 2
cannot cleave effector caspases directly.
The involvement of caspase-2 in the execution of oocyte apoptosis has been
demonstrated by Bergeron et al. (1998). Caspase 2 deficient mice are viable but the
ovaries in 3-4 day females display an excess number of germ cells, and the oocytes are
resistant to chemotherapeutic drug-induced apoptosis. A study on mouse fetal oocytes
cultured in conditions allowing meiotic prophase I progression (Lobascio et al. 2007)
reported an earlier expression of caspase 2. Procaspase 2 is the only procaspase present
constitutively in the nucleus suggesting possible direct DNA DSB-induced caspase 2
activation. Hanoux et al. (2007) showed that caspase-2 constitutes an early step in
triggering the mitochondrial apoptotic pathway in irradiated quiescent oocytes and is
activated before caspase 9 and 3. However, the role of caspase 2 in a specific apoptotic
pathway is controversial.
Another possible player in the apoptosis of oocytes is caspase 9. Prenatal
oocytes in cultured ovaries from caspase 9 -/- mice are more resistant to apoptosis
induced by cytokine deprivation (Maravei et al. 2001). Irradiation also induces cleavage
of caspase 9 in irradiated oocytes at 1 dpp (Hanoux et al. 2007). Caspase 9 and 8 activity
was detected in ovulated oocytes (Papandile et al. 2004).
In contrast, no effector caspases are known as possible players in oocyte
apoptosis before follicle formation. Studies on caspase 3 knockout mice suggested that it
is dispensable for germ cell apoptosis in the female (Matikainen et al. 2001) but
functionally required for granulosa cell apoptosis. Caspase 3 is activated in irradiated
quiescent primary oocytes (Hanoux et al. 2007) and ovulated oocytes (Papandile et al.
2004) but is absent in cultured fetal oocytes (Lobascio et al. 2007). Caspase 6 is
upregulated in female primordial germ cells at 13.5 and 14.5 dpc (Runyan et al. 2006)
but in cultured fetal oocytes caspase 6 remained undetectable (Lobascio et al. 2007).
1.4.6. Other mechanisms for oocyte elimination
The limited frequency of apoptotic oocytes estimated in different studies
contrasts with the magnitude of the germ cell loss observed during oogenesis (Reynaud
and Driancourt 2000). Estimation of apoptosis using the TUNEL assay in fetal and
21
neonatal mouse ovaries showed a maximum of only 3-5% positive oocytes (De Felici
2005). However, in mammals the loss of germ cells occurring before birth is massive
(Baker 1963). In mouse, the loss of germ cells from 13.5 dpc to birth is approximately
65% (McClellan et al. 2003).
Several hypotheses have been forwarded. It is possible that germ cell apoptosis
may be a very rapid process with the TUNEL positive oocytes being rapidly destroyed.
Some authors (Gondos 1978) have claimed that germ cells may leave the ovary by
traveling through the ovarian epithelium.
1.4.6.1. Caspase independent pathways
It has been suggested that only a part of degenerating oocytes is eliminated by
typical apoptosis and that other forms of programmed cell death are responsible for
eliminating germ cells (De Felici 2005).
Alternative models of programmed cell death (PCD) have been proposed,
including autophagy, paraptosis, mitotic catastrophe, apoptosis-like and necrosis-like
PCD (reviewed in Bröker et al. 2005). Autophagy is characterized by sequestration of
cytoplasm and organells in double or multimembrane autophagic vesicles and which are
degraded by cell lysosomal system. Mitotic catastrophe is triggered by mitotic failure
caused by defective cell cycle checkpoints and the cells may be eliminated in a p53independent manner. DNA damage may trigger the mitotic catastrophe.
These types of cell death are caspase-independent and may be mediated by the
release and activation of noncaspase protease death factors such as cathepsins, calpains
and other proteases. Organelles such as the mitochondria, lysosomes, or ER and plasma
membrane death receptors can be involved in these pathways.
Apoptosis-inducing factor (AIF) is an important factor involved in the
regulation of caspase-independent cell death. A distinct cell-death program appears to be
PARP1 mediated release of AIF (reviewed in Hong et al. 2004). In healthy cells, AIF is
located in the mitochondrial intermembrane space and it is released by a mechanism
distinct from that of cytochrome-c. The release of AIF is regulated by the Bcl-2 family
proteins. AIF translocates, via the cytosol, to the nucleus where it binds to DNA and
provokes caspase-independent chromatin condensation (Candé et al. 2002).
22
Calpains (calcium activated neutral proteases) which participate in various types
of cell death, including both caspase-dependent and caspase-independent pathways
(Bröker et al. 2005), and mTOR, mainly involved in the negative control of autophagy
(Castedo et al. 2002) were found to be involved in oocyte death (Lobascio et al. 2007).
This support the hypothesis that caspase-independent and autophagic pathways may be
activated in oocyte death.
23
1.5. Experimental design
It has been suggested that oocytes with meiotic errors may be eliminated by a
pachytene checkpoint which detects unrepaired DSBs and/or the persistence of repair
intermediates to ensure the quality of the oocyte pool. However, this hypothesis remains
to be supported by experimental evidence. Understanding the mechanism of oocyte
elimination is important for identifying the cause/causes for oocyte loss. There are
studies indicating that apoptosis may be involved in oocyte elimination. However, no
convincing evidence was provided and little is known about the signaling pathway and
molecules involved in oocyte elimination, particularly during meiotic prophase
progression. The purpose of this study is to determine if apoptosis plays a role in oocyte
loss in vivo in fetal and neonatal mouse ovaries during meiotic prophase I. As a model
for meiotic errors we used an Msh5 mutant female mouse strain in which all oocytes are
eliminated by neonatal stages due to a failure in meiotic synapsis and recombination.
1.5.1. Msh5 mutant mouse
MSH5 (Mutant S homologue 5) belongs to the Mut S protein family with a role
in mismatch DNA repair. However, MSH5 does not have functions in mismatch repair
but has a role in meiosis (Hollingsworth et al. 1995, Edelmann et al. 1999, de Vries et al.
1999). In a study using purified hMSH4 and hMSH5, Snowden et al. (2004)
demonstrated that these proteins form a heterodimer which recognizes and forms a
sliding clamp at the site of initial cross-over on the pro-Holliday Junction, an early
intermediate of Holliday Junction. They proposed that this heterodimer stabilizes and
preserves meiotic bimolecular double-strand break repair intermediates. It appears that
the MSH4-MSH5 dimer would drive recombination intermediates toward a crossover,
rather than a non-crossover pathway.
It has been shown that the expression of Msh5 in male mouse gonads coincides
with the onset of meiosis (Edelmann et al. 1999). Both male and female mice deficient in
Msh5 are infertile. In the mutant male, spermatogenesis is disrupted. Meiosis does not
progress beyond the zygotene stage, often containing asynapsed chromosomes with
univalents. The condensation of chromosomes corresponds to either the zygotene or
24
pachytene stage (Edelmann et al. 1999, de Vries et al. 1999). It was suggested (de Vries
et al. 1999) that a pseudo G2/M checkpoint mechanism may eliminate defective meiotic
cells. A similar phenotype was observed in mutant females in which most of the oocytes
were blocked at the stage of SC formation at the zygotene-pachytene transition (de Vries
et al. 1999). It has been speculated that the failure of homologous chromosome pairing
may trigger an apoptotic checkpoint which will result in complete ovarian degeneration.
Msh5 mutant females contain fewer oocytes compared to the wild type at 3 dpp and lack
discernible ovaries at puberty.
A study on Spo11/Msh5 and Msh5/Mlh1 double mutants showed that Spo 11
(encoding a protein with a role in initiating DSBs during leptotene) is epistatic to Msh5
and Msh5 is epistatic to Mlh 1 (encoding a protein with a role in late stages of meiotic
recombination). These results suggest that the severe phenotype in Msh5 mutant mice is
caused by defects in DSBs repair (Di Giacomo 2005).
1.5.2. Aim 1
Changes in the number of germ cells in fetal and neonatal ovaries
We studied the number of germ cells in MT (Msh5-/-) comparing to WT
(Msh5+/+) / HT (Msh5+/-) during ovarian development to determine at which stage of
meiotic prophase MT ovaries begin to lose their oocytes due to MSH5 deficiency. In
particular, we wanted to see if a significant loss of germ cells in MT is associated with
the pachytene stage. Experimentally, we first examined the relative abundance of germ
cells in fetal and neonatal ovaries at 14.5-22.5 dpc by immunolabelling of ovarian
sections with antibodies for specific germ cell markers: germ cell nuclear antigen 1
(GCNA1) and mouse vasa homologue (MVH). Then, for a more accurate estimation of
total number of germ cells in ovaries, we prepared dissociated cells spread on slides from
the whole ovaries at 14.5, 16.5, 17.5, 18.5, 19.5, and 20.5 dpc. Germ cells were double
immunolabeled for GCNA1 and synaptonemal complex components (SC) and the total
number of GCNA1-positive cells per ovary was counted.
25
1.5.3. Aim 2
Analysis of meiotic prophase progression
Meiotic progression in WT/HT and MT was examined in dissociated cell
preparations with immunolabelling for SC. For better identification of meiotic stages in
WT as well as the meiotic defect in MT we introduced a ɣH2AX staining. H2AX is a
minor isoform of histone H2A in mammals. This histone is the direct target of ATM
kinase in response to DNA damage. H2AX is phosphorylated at the carboxy-terminal
serine 139 (ɣH2AX) immediately after formation of DSBs by Spo11 during meiotic
prophase (Mahadevaiah et al. 2001). The disappearance of ɣH2AX is correlated with
synapsis at pachytene.
1.5.4. Aim 3
Study of apoptosis
We first performed a TUNEL assay on ovarian sections to see if DNA
fragmentation due to apoptosis occured in WT and MT ovaries. However, this method
did not allow the identification the TUNEL-positive cell type.
To capture and identify the apoptotic pathway, we examined the cleavage of various
caspases and PARP1 in WT and MT ovaries by western blot analysis. If the loss of germ
cells is due to meiotic errors during normal development we anticipated that the intrinsic
mitochondrial pathway would be activated. We also anticipated a higher level of
activation of the same pathway in MT. We chose to investigate the activation of caspase
9, the initiator caspase in the mitochondrial pathway. We also studied the activation of
effector caspase 3, 6, and 7. Because it was reported that caspase 2 is involved in oocyte
apoptosis (Bergeron et al. 1998, Hanoux et al. 2007) we included this molecule in our
study. We also studied the cleavage of PARP1 as a late marker of apoptosis.
The detection of proteins in the whole ovary by western blot does not
discriminate the events in oocytes from those in somatic cells. Therefore, we performed
immunolocalization experiments of cleaved caspases or PARP1 on ovarian sections to
confirm if the apoptotic pathway is activated in oocytes.
26
2. Materials and Methods
2.1. Animals
All animal experiments were conducted in accordance with the Guide to the Care
and Use of Experimental Animals issued by the Canadian Council on Animal Care and
with the approval from the Animal Research Committee of McGill University. Starting
from two Msh5 +/- males, a gift from Dr. Edelmann (Department of Cell Biology, Albert
Einstein College of Medicine, New York) we propagated our mutant colony on the B6
background. The B6 females were purchased from the Jackson Laboratory. The Msh5 +/males were crossed with B6 females and the resulting heterozyous male and female
offspring were crossed to produce Msh5 +/+ (WT), Msh5 +/- (HT) and Msh5 -/- (MT).
The gestation age was defined as days postcoitum (dpc) assuming that the copulation
occurred at 1:00 AM. Although delivery occurs usually at 19.5 dpc we used the same
method for counting the ages of newborn mice.
2.2. Genotyping
Genotyping was performed by PCR analysis of ear punch lysis for adult mice or
tail lysis for embryos and newborn mice.
2.2.1. Separate amplification of wild-type and mutant alleles
Primers A5 and C5 amplify a 300-bp fragment from the wild-type allele; primers
A5 and B amplify 400-bp fragment from the mutant allele. The thermo-cycling
conditions were: 94ºC 10min; 45 cycles of 94ºC 1min, 55ºC 1 min and 72ºC 1 min
followed by extension at 72ºC 5 min.
Primer A5
AGACCTGCACTGCGAGATCCG
Primer B
TGGAAGGATTGGAGCTACGG
Primer C5
TCTCGAATAGCCGTAGTCCCG
2.2.2. Duplex PCR
The primers used in the first method did not allow amplification of both alleles in a
single reaction. Therefore the primers were redesigned as follow:
27
Primer A’
AGGAGCCCGTGGTAGGAG
Primer B
TGGAAGGATTGGAGCTACGG
Primer C’
CCATGGATACAGGGAGAGTAATG
Primers A’ and C’ amplify a 300-bp fragment from the wild-type allele; primers A’
and B amplify a fragment at around 400-bp from the mutant allele. The cycling
conditions were: 95ºC 2 min; 35 cycles of 95 ºC 30 s, 55 ºC 30s and 72 ºC 1 min
followed by final extension at 72 ºC 15 min.
In some experiments the genotype was confirmed by using purified DNA.
2.3. Isolation of ovaries
Pregnant mice were sacrificed and their foetuses were removed at 14.5-18.5 dpc.
Neonatal mice were sacrificed at 19.5, 20.5 and 22.5 dpc and their mothers kept for the
next experiments. Ovaries with or without mesonephroi were removed under a dissecting
microscope and kept in Eagle’s minimal essential medium (MEM) with Hanks’
supplemented with 0.25 mM Hepes buffer (Invitrogen) until further processing.
2.4. Fixation of ovaries for histological preparation
Isolated ovaries were fixed in 2% formaldehyde in microtubule-stabilizing buffer
(Mesinger and Albertini 1991) for 1 hour at room temperature. After fixation, ovaries
were washed in PBS and kept in 70% ethanol at 4ºC until embedding.
2.5. Paraffin embedding and microtome sectioning of fixed ovaries
Ovaries were dehydrated in 95% ethanol, two changes of absolute ethanol and one
change of toluene for 45 minutes each at room temperature. They were placed in a
second bath of toluene for 30 minutes and transferred into an oven at 60ºC for 15
minutes. The toluene was removed and ovaries were incubated two times in melted
paraffin at 60ºC for 45 minutes each. Ovaries were removed from paraffin and placed in
metal molds containing paraffin on a hot plate at 62ºC. On top of each mold was placed
an embedding ring. The molds were placed on ice and filled with hot paraffin and left to
room temperature until solidified. After solidification of paraffin the molds were left on
28
ice water and the embedding rings removed. The paraffin blocks were kept at 4ºC until
sectioning.
Ovaries were sectioned at 6 µm and serial sections from each ovary were placed on
histology slides. Slides were incubated in an oven at 60ºC for 20 minutes, transferred in
glass jars with silica gel and kept at 4 ºC.
2.6. Deparaffinization and hydration
Slides were immersed in three changes of toluene for 2 minutes each, one change
of absolute ethanol, two changes of 95% ethanol, one change of 70% ethanol and rinsed
in ddH2O.
2.7. Antigen retrieval
After deparafinization, slides with histological sections were kept in Tris-buffer
saline 0.05 M (pH 7-8). A styrofoam bath with water was heated in a microwave until
boiling. Beakers containing 0.05 M Tris-HCl (pH=10.0) were placed in the stylo-form
bath and the slides were immediately transferred to the beakers, covered and left for 30
minutes. The beakers were then cooled down in running water. Slides were washed
briefly in distilled water.
2.8. Dissociated cells and chromosome spread preparations on histological
slides by cytospin centrifugation
The ovaries were isolated from the adjacent mesonephroi and surrounding tissues
in MEM (H). One ovary was transferred with a small volume of MEM (H) into a 1.5 ml
microfuge tube. The medium was removed as much as possible with a Pasteur pipette
and replaced with 100 µl of 0.05% collagenase in MEM (H) followed by an incubation
for 15 minutes at 37 ºC. Subsequently, the ovaries were rinsed with 100 µl MEM (H) and
incubated with 100 µl of 0.125 % trypsin in Rinaldini solution (Rinaldini 1959) at 37 ºC
for 12 minutes. The action of trypsin was stopped by adding to each tube 10 µl of 2%
soybean trypsin inhibitor (Worthington) followed by gentle mixing. The solution was
gently removed and replaced with 100 µl of 10 % of FBS in MEM (H). The ovaries were
rinsed twice with 100 µl MEM (H) and once with 100 µl of Ca2+, Mg2+-free Dulbecco’s
29
PBS (DBPS). Finally the ovaries were dissociated in 100 µl of Ca2+, Mg2+-DBPS by
gentle pipetting and centrifuged at 360 g for 3 minutes at room temperature. The
supernatant was gently removed and the pellet was resuspended in 20 µl of MEM (H).
The cell suspension was applied to each well of cytospin centrifuge chamber (Thermo
IEC, Needham Heights, MA) containing 400 µl of 0.5 % NaCl solution (pH 8.0) placed
on a histology slide, and incubated for 5 minutes. The cells were spun down onto the
slide, followed by two fixations in 400 µl of 1% paraformaldehyde solution (pH 8.2) and
three washes in 0.4% Photoflo (Kodak, Eastman, NY) in H2O (pH 8.0). After each step
slides were spun down. Slides were then vacuum dried and stored in sealed boxes with
silica gel at -20 ºC.
2.9. TdT-mediated dUTP nick end labelling (TUNEL)
After deparaffinization, ovarian sections were incubated with 20 µg/ ml proteinase
K in 0.5 M Tris-HCl (pH=7.5) for 15 minutes at 37ºC in a water bath. Slides were then
rinsed three time in PBS with stirring and incubated with 50 µl TUNEL reaction mix (In
Situ Cell Death Detection Kit, Roche) for 60 minutes covered with parafilm in a humid
chamber at 37 ºC. Slides were then flushed with PBS, washed three times each for 10
minutes in PBS with stirring and rinsed twice in double distilled water. After being dried
in vacuum, slides were mounted in Prolong Antifade with 4’,6-diamidino-2-phenylindole
(DAPI) and left at least 20 minutes in dark at the room temperature before microscope
observations.
2.10. GCNA1 and MVH immunolabelling of ovarian sections
After deparaffinization and hydration, slides were incubated three times each for
10 minutes in holding buffer (HB) containing 1% goat serum, 0.3% bovine serum
albumin, and 0.005% triton X-100 in PBS (Dobson et al. 1994). In some experiments,
we used antigen retrieval before washings, as described. After washing slides were
incubated with rat monoclonal anti-GCNA1 or rabbit polyclonal anti-MVH antibody for
1 hour in a humidified chamber at room temperature. Then, slides were rinsed with PBS,
washed three times each for 10 minutes in HB and incubated with a secondary antibody
for 45 minutes in a humidified chamber at room temperature. The details on the primary
30
and secondary antibodies are given in Table 1 and 2. The slides for GCNA1
immunolabelling were rinsed with PBS, washed three times each for 10 minutes in HB
and incubated with avidin–FITC at 1:1000 dilution in a humidified chamber in the dark
at room temperature. After incubation slides were rinsed with PBS, washed three times
each for 10 minutes in PBS, rinsed twice in distilled water, dried under vacuum and
mounted with Prolong Antifade Kit (Invitrogen) with 0.4 µg/ml DAPI.
2.11. Double immunolabelling of GCNA1 and ɣH2AX of ovarian sections
After antigen retrieval the procedures were similar to those for GCNA1
immunolabelling. Slides were incubated with the primary antibodies overnight in a
humid chamber at room temperature. The details on the primary antibodies, secondary
antibodies and fluorescent-labelled solution are given in Table 1 and 2.
.
2.12. Double imunolabelling of GCNA1 and cleaved Caspase 9 (37/15 KDa or
39 KDa) or cleaved PARP1 (24KDa) of ovarian sections
After antigen retrieval the procedures were similar to those for GCNA1 and
ɣH2AX immunolabelling. A blocking step with avidin-biotin was introduced before the
incubation with the primary antibodies. The details on the primary antibodies, secondary
antibodies and fluorescent-labelled solution are given in Table 1 and 2.
2.13.
Immunocytochemical
staining
of
GCNA1/SC,
SC/ɣH2AX
and
GCNA1/SC/ɣH2AX of chromosome spreads
The protocol followed the same steps as for double immunolabelling of ovarian
sections with GCNA1 and ɣH2AX immunolabelling without antigen retrieval. The
details on the primary antibody, secondary antibody and fluorescent-labelled solution are
given in Table 1 and 2.
31
Table 1. Reagents used for immunofluorescent staining
Type of staining
Sample
Primary antibody
GCNA1
Histological rat monoclonal anti-GCNA1 1:20
sections
MVH
Histological rabbit polyclonal anti-MVH 1:1000
sections
GCNA1/ɣH2AX
Histological rat monoclonal anti-GCNA1 1:10
sections
rabbit polyclonal-anti ɣH2AX 1:400
GCNA1/cleaved
Histological rat monoclonal anti-GCNA1 1:10
Caspase 7 (20 KDa) sections
rabbit polyclonal anti-cleaved caspase
7 (20 KDa) 1:50, 1:100, 1:300
GCNA1/cleaved
Histological rat monoclonal anti-GCNA1 1:10
Caspase 9 (35/15 sections
rabbit polyclonal anti-cleaved caspase
KDa)
9 (35/15 KDa) 1:500, 1: 1000
GCNA1/cleaved
Histological rat monoclonal anti-GCNA1 1:10
Caspase 9 (37 KDa) sections
rabbit polyclonal anti-cleaved caspase
9 (37 KDa) 1:50
GCNA1/cleaved
Histological rat monoclonal anti-GCNA1 1:10
PARP1 (24KDa)
sections
rabbit monoclonal anti-cleaved
PARP1 (24 KDa)
1:50, 1:100, 1:300, 1:400
GCNA1/SC
Chromosome rat monoclonal anti-GCNA1 1:10 or
spread
1:20
rabbit polyclonal anti-SC 1:1000
SC/ ɣH2AX
Chromosome mouse polyclonal anti-SC 1:1000
spread
rabbit polyclonal anti-ɣH2AX 1:2000
GCNA1/SC/ɣH2AX Chromosome rat monoclonal anti-GCNA1 1:1
spread
mouse polyclonal anti-SC 1:1000
rabbit polyclonal anti-ɣH2AX 1:2000
Secondary antibody
goat anti-rat biotin 1:1000
Avidin-conjugate
avidin–FITC 1:1000
goat anti-rabbit rhodamine 1:1000
goat anti-rat biotin 1:1000
avidin–FITC 1:1000
goat anti-rabbit rhodamine 1:1000
goat anti-rat FITC 1:1000
streptavidin-rhodamin
goat anti-rabbit biotin 1:1000
1:1000
goat anti-rat FITC 1:1000
goat anti-rabbit biotin 1:1000
streptavidin-rhodamin
1:1000
goat anti-rat FITC 1:1000
goat anti-rabbit biotin 1:1000 or
1:200
goat anti-rat FITC 1:1000
goat anti-rabbit biotin 1:1000
streptavidin-rhodamin
1:1000
goat-anti-rat FITC 1:1000
goat anti-rabbit biotin 1:1000
avidin-Cy3 1:1000
goat anti-mouse biotin 1:1000
goat anti-rabbit FITC 1:1000
goat anti-rat Alexa (350) 1:25 or
1:50
goat anti-mouse biotin 1:1000
goat anti-rabbit FITC 1:1000
avidin-Cy3 1:1000
streptavidin-rhodamin
1:1000
avidin-Cy3 1:1000
32
Table 2. The source of the antibodies used for immunofluorescent staining
Antibody
Source
rat monoclonal anti-GCNA1
G. C. Enders
rabbit polyclonal anti-MVH
T. Noce
rabbit polyclonal anti-ɣH2AX
Abcam
rabbit polyclonal anti-cleaved caspase 7
Cell Signaling
rabbit polyclonal anti-cleaved caspase 9 (35/15
KDa)
Imgenex
rabbit polyclonal anti-cleaved caspase 9 (37 KDa)
Cell Signaling
rabbit monoclonal anti-cleaved PARP1 (24 KDa)
Abcam
rabbit polyclonal anti-SC
P. Moens
mouse polyclonal anti-SC
P. Moens
goat anti-rat biotin
PIERCE
goat anti-rabbit biotin
PIERCE; VECTOR
goat anti-mouse biotin
PIERCE
goat anti-rat FITC
PIERCE
goat anti-rabbit FITC
VECTOR
goat anti-rabbit rhodamine
PIERCE
goat anti-rat Alexa
INVITROGEN
avidin–FITC
VECTOR
streptavidin-rhodamin
INVITROGEN
avidin-Cy3
PIERCE
33
2.14. Counting and staging of germ cells
The total number of germ cells was counted for each ovary under the light
microscope. In most samples around 100 cells/sample were analysed for meiotic stages.
However, in some samples, especially at advanced developmental stages when the total
number of oocytes is low, fewer cells were staged.
2.15. Statistical analysis
Student’s t test was used to compare the results of germ cell counting among the
three genotypes of Msh5. The s.e.m. was calculated for each group.
2. 16. Western Blot
Ovaries at 16.5, 17.5, 18.5, 19.5 and 21.5 dpc were collected in MEM (H) and snap
frozen in liquid nitrogen or they were stored in sample buffer and kept at -80 ºC until
use. Two ovaries were pooled, unless specified, and processed in 15 or 20 µl sample
buffer (Tris 50 mM, SDS 2%, bromphenol blue 0.1%, Ficoll 2.5%, dithiotreitol 100 mM)
with protease inhibitor cocktail (Roche) by repeated freezing and thawing and
dissociated by vortexing followed by centrifugation. After dissociation the samples were
boiled for ten minutes. The sample lysate was loaded on a 14-16% polyacrylamide gel
(PAG) containing SDS and the electrophoresis was run at 25 mA. A 4-15 % gradient gel
(Biorad) was also used. Equal amounts were loaded for paired samples (WT and MT) of
the same age. Molecular weight standards (kaleidoscope plus protein molecular standard
(BioRad), biotinylated molecular weight standards) and control proteins (Jurkat Cell
Lysates) were also applied. Proteins in the SDS-PAGE were electro transferred to a
nitrocellulose membrane in Tris-glycine buffer containing 20% methanol. After transfer
the membranes were blocked in 5% skim milk in PBST for 1 hour and incubated with
the primary antibody diluted 1:1000 in PBST overnight at 4ºC. We used rabbit
polyclonal primary antibodies recognizing the full forms as well as cleaved forms or
only the cleaved form of different caspases and PARP 1 (Table 3). Next day the
membranes were washed in PBST and incubated for 2 hours with donkey anti-rabbit
conjugated with biotin at 1:5000 dilution in PBST. The membranes were washed and
incubated for 30 minutes with a goat-anti biotin conjugated with horse radish peroxidase
34
(Cell Signaling) at 1:500 dilution. The enzymatic activity was detected with a LumiLight ECL kit (Roche). After detection the membranes were stripped and reblotted for
other antibodies. If stripping was not performed immediately the membranes were kept
at -4 ºC or frozen in PBS. For stripping we used a ReBlot Plus Mild Antibody stripping
solution (Millipore) 1X prepared according manufacturer indications. The membranes
were incubated for 15 minutes in 1X stripping solution with gentle agitation and washed
three times 5 minutes each in PBST. The efficiency of stripping was tested by detection
with Lumi-Light ECL kit.
2. 17. Specificity of antibodies
2. 17. 1. Blocking peptide
The specificity of Caspase 2L was verified using a blocking peptide for Caspase
2L (Santa Cruz). Homogenous samples were loaded on the same gel and subjected to
western blot. After transfer the membrane was cut in two and subjected to incubation
with caspase 2L antibody 1:1000 dilution or with caspase 2L 1:1000 dilution treated in
advance with blocking peptide (caspase 2L : blocking peptide 1:15). The bands
corresponding to 51 KDa and 12-13 KDa did not show when the mixture of caspase 2
and blocking peptide was used (Fig. 16).
2. 17. 2. Jurkat Apoptosis Cell Lysates
We used Caspase 3 Jurkat Apoptosis Cell Lysates from Cell Signaling
Technology. The cytoplasmic fraction obtained from untreated Jurkat cells lysed in
Chaps cell extract serves as a negative control while the extract treated with cytochrome
c in vitro serves as a positive control for caspase 3 cleavage.
We also used Jurkat Cell Lysates (Cell Signalling) untreated (negative control) or
treated with 25 µm etoposide (positive control) for the induction of apoptosis. The
treatment with etoposide activates an apoptotic cascade and induces proteolytic cleavage
of various apoptotic-related proteins, including caspases and PARP1. We used this lysate
pair to confirm the molecular weights for full forms and cleaved forms of caspases 3, 6,
7 and PARP1.
35
2. 18. Analysis of band intensity
The bands corresponding to the cleaved and uncleaved forms were quantified using
UN SCAN IT software and the cleaved/full form ratios were calculated and used for
comparison. When more than one experiment was performed the mean ratio of cleaved/
uncleaved form was used for comparison.
Table 3. Specification of the antibodies used for western-blot
Antibody
Source
Antigen
Immunized
MW
Caspase 2L
Santa Cruz
mouse
Rabbit
12, 13, 51
Caspase 3
Cell Signaling
human
Rabbit
17, 35
Caspase 6
Cell Signaling
unspecified
Rabbit
15, 35
Caspase 7
Cell Signaling
human
Rabbit
20, 35
Caspase 9
Cell Signaling
mouse
Rabbit
37, 39, 49
Cleaved
Cell Signaling
mouse
Rabbit
37
Cleaved
caspase 9
PARP1
Imgenex
human
Rabbit
15, 35
Cell Signaling
unspecified
Rabbit
24, 89, 116
Cleaved
Cell Signaling
mouse
Rabbit
89
Caspase 9
PARP1
Bold - full form
36
3. Results
3.1 Changes in the number of germ cells in fetal and neonatal ovaries
3.1.1 The abundance of germ cells in ovarian sections detected by GCNA1 and
MVH immunolabelling
To observe the difference in the abundance of the germ cells between the MT and
the WT/HT ovaries at 14 – 22.5 dpc we used immunolabelling of ovarian sections with
the antibodies against germ-cell nuclear antigen 1 (GCNA1) or mouse vasa homologue
(MVH). GCNA1 is known to be expressed in the germ cells in fetal and neonatal ovaries
(Enders and May 1994). MVH is expressed in the cytoplasm of germ cells until late
postnatal life (Toyooka et al. 2000). In some experiments we used a double
immunolabelling of GCNA1/ ɣH2AX and the results on GCNA1 immunolabelling were
included here. For most of the cases we used WT and MT ovaries for comparison. When
WT was not available HT was used for comparison since we did not observe major
differences between them. The details on the number of WT/MT pairs of ovaries are
given in Table 4.
Table 4. The number of WT/MT pairs of ovaries investigated using different methods
Age (dpc)
GCNA1
GCNA1/ ɣH2AX
MVH
14.5
1
0
1
16.5
1
3
1
17.5
0
2
0
18.5
2
1
1
19.5
0
1
0
20.5
2
2
2
22.5
1
1
3
We did not observe major differences in the abundance of GCNA1 positive cells
between MT and WT until 19.5 dpc (Fig. 4). However at 18.5 dpc we observed a higher
variability in germ cell abundance in MT. At 20.5 dpc there was a higher abundance of
37
oocytes in WT comparing to MT (Fig. 4 M, O). At 22.5 dpc very few oocytes were
observed in MT (Fig. 4 S).
The results were consistent with the cytoplasmic immunolabelling with MVH. No
major difference was observed between MT and WT up to 20.5 dpc (Fig. 4). At 20.5 dpc
the WT showed large cytoplasmic staining while in MT the stain was very weak or
absent (Fig. 4 N, P). At 22.5 dpc the MVH staining was more intense in the WT (Fig. 4
R) compared to the anterior stages due to an increase in oocyte size while in MT this
staining was absent (Fig. 4 T).
38
Figure 4. Immunolabelling of ovarian sections for GCNA1 (green) or MVH (red).
Blue color represents DAPI. The yellow staining represents blood cells. The bar
indicates 200 µm.
A.14. 5 dpc WT ovary imunolabeled for GCNA1
B. 14.5 dpc WT ovary imunolabeled for MVH
C. 14. 5 dpc MT ovary imunolabeled for GCNA1
D. 14.5 dpc MT ovary imunolabeled for MVH
E. 16. 5 dpc WT ovary imunolabeled for GCNA1
F. 16.5 dpc WT ovary imunolabeled for MVH
G. 16. 5 dpc MT ovary imunolabeled for GCNA1
H. 16.5 dpc MT ovary imunolabeled for MVH
I. 18. 5 dpc WT ovary imunolabeled for GCNA1
J. 18.5 dpc WT ovary imunolabeled for MVH
K.18. 5 dpc MT ovary imunolabeled for GCNA1
L. 18.5 dpc MT ovary imunolabeled for MVH
M. 20. 5 dpc WT ovary imunolabeled for GCNA1
N. 20.5 dpc WT ovary imunolabeled for MVH
O. 20. 5 dpc MT ovary imunolabeled for GCNA1
P. 20.5 dpc MT ovary imunolabeled for MVH
Q. 22.5 dpc WT ovary imunolabeled for GCNA1
R. 22. 5 dpc MT ovary imunolabeled for MVH
S. 22.5 dpc WT ovary imunolabeled for GCNA1
T. 22.5 dpc MT ovary imunolabeled for MVH
39
A
B
E
I
C
F
G
J
M
Q
D
H
K
N
R
O
L
P
S
T
40
3.1.2. Changes in the number of GCNA1 positive germ cells examined in spread cell
preparations
For a more accurate estimation of the total number of germ cells in ovaries, we
prepared cell spread slides from the whole ovary (14.5, 16.5, 17.5, 18.5, 19.5, 20.5 dpc).
The total number of GCNA1 positive cells/slide was counted. A gradual loss of germ
cells was observed in the WT, HT and MT from day 16.5 to 20.5 dpc (Fig. 5). The
number of germ cells was consistently lower in MT. A significant decrease in MT
comparing to the WT (p<0.05) was found at 19.5 dpc and in MT comparing to the HT at
20.5 dpc.
Figure 5. Changes in the total number of GCNA1 positive cells from ovaries. The
number of ovaries are indicated on the top of each value.
41
3.2. Meiotic prophase progression in WT, HT and MT ovaries
3.2.1. Identification of meiotic stages using SC/GCNA staining
The first analysis of meiotic stages was based on the staining of SC with a
polyclonal antibody recognising both SCP1 and SCP3. SCP3 starts to form at L and
disappears at the end of D. SCP1 starts to form in Z and disappears in D before SCP3
disappears. Therefore, we observed the localization of SCP3 with the antibody used. The
criteria for identifying the meiotic stages are given in Tables 5 and 6 and the results on
meiotic progression using GCNA1/SC labelling are shown in the Fig. 6 and 7. The
results from WT and HT ovaries were pooled since we did not observe major differences
between them. At all examined stages in WT/HT average proportion of Z was high. The
proportion of P was relatively constant. However, at stage 17.5 dpc we observed a very
low proportion of P and a high proportion of L in three samples out of four but this may
be due to experimental conditions in one experiment. In this experiment the SC staining
was very weak and it is possible that the SC proteins were not well preserved. The
proportion of D was low at 16.5-18.5 dpc and increased at 19.5 dpc.
With this method we found it difficult to distinguish between Z and D stages in
WT/HT since the appearances of SCP1 and SCP3 were similar between early Z and late
D or between late Z and early D. While some meiotic configurations showed a typical
appearance of Z or D others were hard to assign to a category. In literature there are
some additional criteria used to define meiotic stages such as the bouquet configuration
characteristic for Z or the higher volume of oocytes in D compared to other stages.
However, due to the use of hypotonic treatment these criteria were not very useful. The
bouquet configuration may be lost in some cells. The hypotonic treatment may also
affect the volume of nuclei. In histological preparations oocytes in a certain stage of
meiotic prophase may be clustered. However this is not the case in spread cells. Another
difficulty is P staging which is defined as the completely synapsed stage and is often
assumed to have condensed SC. However there are early P configurations with very long
and tangled SC which may be misclassified as Z.
Identifying meiotic stages in the MT was also difficult. While some of oocytes
with condensed SC and incomplete synapsis may be identified as abnormal P or
42
abnormal Z, the stage of oocytes at 19.5 dpc was ambiguous. No normal P was observed
in MT.
3.2.2. Identification of meiotic stages using SC/ ɣH2AX staining
For a better delimitation of the zygotene and diplotene stages we introduced a double
immunolabelling of SC and ɣH2AX. The results of the triple GCNA1/SC/ɣH2AX
staining were also included here. The criteria used to identify the meiotic stages in
WT/HT and MT with this method are given in Table 5 and Table 6. The SC
characterization is the same as for GCNA1/SC staining. In normal meiosis (WT) ɣH2AX
appears in L, it is extensively present in early Z and starts to disappear with the
formation of synapsis and DNA repair (Fig. 10). At P and D ɣH2AX is negative.
However positive ɣH2AX residues may persist along SC in some oocytes at P. Some
positive spots (1-3) may be seen in some P oocytes, probably corresponding to
incomplete synapsis. Occasionally, in WT some cells with the SC characteristic for D
show regions of positive ɣH2AX but these are smaller compared to a similar Z stage.
In MT we defined a special category as Z-P abnormal (Z-P-ab). This category
includes normal appearing Z and abnormal P since it is difficult to establish a clear
delimitation between these two stages. However, this category describes different
subcategories in different stages of ovarian development as follows:
- At 16.5 dpc most of the oocytes may be classified as Z. Some cells may be
classified as abnormal Z because the number of SC appears to be doubled (Fig. 11 a).
This may be due to the failure of asynapsis. A small percentage of cells may be classified
as P abnormal.
- At 17.5, 18.5 and 19.5 dpc the synaptonemal complex becomes more condensed.
Most of the oocytes look as late Z – P abnormal- early D with different patterns of
positive ɣH2AX (Fig. 11 b-f). The SC is condensed as in P but with extensive asynapsis
or completely asynapsed chromosomes.
- At 20.5 dpc SC is decondensed and asynapsed and or /discontinuous. The
category of P abnormal with condensed SC is missing. ɣH2AX is present as positive
areas which in some cases are smaller when comparing to Z cells with the same degree
43
of SC condensation (Fig. 11 i). Therefore these cells may be also classified as D
abnormal.
In MT, we classified those cells with desynapsed/incomplete/decondensed SC and
ɣH2AX negative as D (Fig. 11 g, h, j).
The results using SC/ ɣH2AX technique are shown in Fig. 8 and 9. In WT meiosis
progressed gradually showing a peak of pachytene at 17.5 dpc. At 18.5 – 19.5 dpc the
predominant stage was D and by 20.5 dpc all cells entered D. The average proportion of
Z was smaller but the average proportion of D is higher from 17.5 to 19.5 dpc comparing
to the previous technique. Other differences such as a higher abundance of P at 16.5 dpc
and 17.5 dpc in SC/ ɣH2AX labelled samples or a higher proportion of L in GCNA1/SC
labelled samples may be explained by the fact that different populations of cells were
analyzed by the two methods. In the GCNA1/SC technique we identified the meiotic
stages of GCNA1 positive cells while in the second technique we staged oocytes positive
for SC. During the first observations we classified as P mostly the oocytes with complete
synapsis and more condensed SC. Using ɣH2AX technique we considered as P those
cells with SC completely formed and ɣH2AX negative but with different degree of SC
condensation. Cells with minor desynapsis and ɣH2AX negative or only 1-2 minor spots
were also considered as P.
In MT, as with the previous technique, no normal P was observed at any
developmental stage. Univalent-like chromosomes were also observed (Fig. 11 e). In
these oocytes, SC was short, condensed and thick. The thickness of the SC in this case is
more characteristic of normal P.
These results suggest that meiosis in MT is blocked at Z-P transition due to
persistence of unrepaired DSBs and asynapsis.
44
Table 5. Criteria used for defining meiotic stages in WT
Meiotic stage
SC / (GCNA1)
(SC) / ɣH2AX
Leptotene (L)
SC appears
Some positive spots
Zygotene (Z)
SC is more advanced
Uniform and bright (early Z),
(Fig. 10 b, c, d)
than
patched (late Z) or pale (Z-P
(Fig. 10 a)
in
L
incompletely
but
formed
transition)
and/or asynapsed
Pachytene (P)
SC
is
completely
(Fig. 10 e, f)
formed and synapsed
Negative. Some positive spots (13) or ɣH2AX residues along SC
may be observed in some oocytes
Diplotene (D)
SC is desynapsed
Negative
(Fig. 10 g, h, i)
and/or incomplete. In
late D the SC may look
like early Z or L
Complex
GCNA 1 positive but
This staining did not allow the
negative
SC negative. It may
detection of this category
correspond
to
germ
cells which did not
enter L or those in late
D
in
which
SC
disappeared.
45
Table 6. Criteria used for defining meiotic stages in the MT
Meiotic stage
SC/(GCNA1)
SC/ ɣH2AX
Leptotene (L)
SC starts to form
Some positive spots
Zygotene-Pachytene
Zygotene: SC is more advanced Positive:
abnormal
bright,
(Z-Pab) than in L but incompletely patched, one positive
(Fig. 11 a-f, i)
formed and/or asynapsed.
Pachytene
abnormal:
area or intense traces
SC along SC
condensed as in P but with
extensive
asynapsis
completely
or
asynapsed
(univalents).
These 2 categories have been
pooled.
Pachytene normal (P)
SC
completely
formed
and Negative.
synapsed. Extremely rare in MT
Diplotene (D) (Fig. SC is desynapsed and/or
11 g, h, j)
incomplete
Complex negative
GCNA
1
positive
but
Extremely
rare in MT
Negative
SC This technique did not
negative. It may correspond to allow the detection of
germ cells which did not enter L this category
or those in late D in which SC
disappeared
46
Figure 6. Meiotic progression during ovarian development in WT/HT identified by
GCNA1/SC immunolabelling. The number of ovaries is indicated on the top of each column.
Figure 7. Meiotic progression during ovarian development in MT identified by GCNA1/SC
immunolabelling. The number of ovaries is indicated on the top of each column.
47
Figure 8. Meiotic progression during ovarian development in WT/HT identified by
SC/ɣH2AX immunolabelling. The number of ovaries is indicated on the top of each column.
Figure 9. Meiotic progression during ovarian development in MT identified by SC/ɣH2AX
immunolabelling. The number of ovaries is indicated on the top of each column.
48
a-L
b - Early Z
f - Mid P
g - Early D
c - Mid Z
d – Late Z - Early P
h - Mid D
e - Early P
i - Late D
Figure 10. Meiotic stages in spread chromosomes from WT ovary double
immunolabeled for SC (red) and ɣH2AX (green). The bar represents 20 µm.
49
a
b
e
c
f
d
g
i
h
j
Figure 11. Meiotic stages in spread chromosomes from MT ovary double
immunolabeled for SC (red) and ɣH2AX (green). The bar represents 20 µm.
a - Z with a double number of SC cores and ɣH2AX intense staining
b, c, d - P-ab with extensive asynapsis and ɣH2AX staining
e - P-ab with univalent SC cores
f - Z-P-ab in which SC appears like D but ɣH2AX positive
g, h – D with ɣH2AX negative
i - oocyte at 20.5 dpc with ɣH2AX positive (Z-P-ab)
j - oocyte at 20.5 dpc with ɣH2AX negative (D)
50
3.2.3. Observation on meiotic progression in ovarian sections
We performed a double GCNA1/ ɣH2AX immunolabelling on ovarian sections to
observe the distribution of oocytes in Z in WT or of oocytes in Z or with persistent
asynapsis in MT. The details on the number of WT/MT pairs of ovaries investigated are
given in Table 4. At 16.5 dpc most of the germ cells were positive for ɣH2AX
suggesting that most of them were in Z or Z-P transition in both genotypes of ovaries
(Fig. 10 a-f). By day 17.5-18.5 dpc the number of positive cells for ɣH2AX was lower in
WT (Fig. 10 g-l). In MT we observed a higher number of ɣH2AX positive cells at 17.5 19.5 dpc comparing to the WT controls. At 20.5 dpc most of the germ cells are negative
for ɣH2AX in both WT and MT. These results show the persistence of unrepaired DSBs
in MT up to 19.5 dpc and confirm the previous observations on meiotic progression.
51
a
b
c
d
e
f
g
h
i
j
k
l
Figure 12. Double immunolabelling of ovarian sections for GCNA1 (green), H2AX
(red) and merged. The bar represents 200 µm.
a-c 16. 5 dpc WT ovary immunolabelled with (a) GCNA1, (b) H2AX and (c)- merged
d - f 16. 5 dpc MT ovary immunolabelled with (d) GCNA1, (e) H2AX and (f)- merged
g - i 18. 5 dpc WT ovary immunolabelled with (g) GCNA1, (h) H2AX and (i)- merged
j – l 18. 5 dpc MT ovary immunolabelled with (j) GCNA1, (k) H2AX and (l)- merged
52
3.3. Apoptosis
3.3.1. TUNEL assay
We used the TUNEL assay of ovarian sections in order to see if DNA
fragmentation due to apoptosis occurs in ovaries of MT and WT. In some cases WT was
replaced with HT for comparison with MT. Two pairs of ovaries were analysed for 16.5,
18.5 and 22.5 dpc, four pairs for 20.5 dpc and one pair for 14.5, 17.5 and 19.5 dpc for
both WT and MT. At 14.5 dpc we observed very few positive cells in both WT and MT
(Fig. 13 a-b). The number of positive cells increased at 16.5, 17.5, 18.5 and 19.5 dpc
(Fig. 13 i-j). By 20.5 dpc the number of positive cells was lower. At 18.5 dpc we
observed a higher number of positive signals in MT compared to WT (Fig. 13 e-f). In
some sections in both WT and MT at 18.5 and 19.5 dpc we observed a high number of
positive signals (up to 60 -70). At a high magnification we observed that some positive
signals were small and may correspond to the somatic cell nuclei (Fig. 13 k) while other
positive cells, judging by the nucleus size and the presence of chromosome-like
structures, may correspond to oocytes (Fig.13 m). Occasionally, TUNEL positive cells
were observed to protrude outside of the epithelium (Fig. 13 l). We concluded that
apoptosis does occur in ovaries of both WT and MT from 16.5 dpc to 19.5 dpc. However
it was difficult to convincingly identify the apoptotic cell type by this method.
53
Figure 13. Distribution of TUNEL positive cells in ovarian sections (green). Blue
represents DAPI and the yellow represents blood cells. The bar represents 200 µm (a-j),
50 µm (k-l) and 20 µm (m).
a - 14. 5 dpc WT ovary
b - 14.5 dpc MT ovary
c - 16.5 dpc WT ovary
d - 16.5 dpc MT ovary
e - 18.5 dpc WT ovary
f - 18.5 dpc MT ovary
g - 19.5 dpc WT ovary
h - 19.5 dpc MT ovary
i - 20.5 dpc WT ovary
j - 20.5 dpc MT ovary
k - Small TUNEL positive signals which may correspond to somatic cells
l - Positive TUNEL cell protrunding from the ovarian surface (arrow)
m - Two TUNEL positive cells showing chromosome-like structures (arrows)
54
a
b
c
d
e
f
g
h
i
j
55
k
l
m
56
3.3.2. Western Blot analysis of the activation of caspase 2, 3, 6, 7, 9 and
cleavage of PARP1
To detect if activation of caspases or cleavage of PARP1 occurs in ovaries in vivo
we used Western Blot analysis.
3.3.2.1. Caspase 9
Caspase 9 is the main initiator caspase of the mitochondrial pathway. Caspase 9
antibody (Cell Signaling) recognizes the full form at 49 KDa and two bands at 39 KDa
and 37 KDa corresponding to the cleaved forms (Fig. 14). The expression of the full
form of caspase 9 and cleaved forms was detected in both WT and MT throughout all
investigated stages. Quantification of 39 KDa in some blots proved to be technically
difficult due to its close proximity to the full form. Since this band had a lower intensity
we quantified the band at 37 KDa. The mean ratio of the cleaved form at 37 KDa to the
full form was calculated as the activation level of caspase 9. The activation of caspase 9
increased during ovarian development reaching a peak at 18.5 dpc and decreased by 19.5
dpc in both WT and MT ovaries (Fig. 14). No difference in activation level was observed
between WT and MT.
3.3.2.2. Cleaved Caspase 9 of 35/15 KDa
The cleavage of Caspase 9 was also studied by using an antibody which detected
the 35 KDa and 15 KDa fragments of caspase 9 (see Table 3). We detected both
fragments (Fig. 15) but the band at 15 KDa was weak. This may be explained by the fact
that the incubation with this antibody was performed after multiple stripping of the
membrane. Since this antibody does not recognise the full form of Caspase 9 we could
not calculate the activation level.
3.3.2.3. Caspase 2 L
Caspase 2L is another potential initiator caspase in the mitochondrial pathway. The
full form of caspase 2L was detected throughout all investigated stages in both WT and
MT (Fig. 17). Caspase 2L showed a similar pattern of activation as caspase 9 however at
much lower levels. The level of activation increased moderately from 16.5 dpc to 18.5
57
dpc and decreased at 19.5 dpc (Fig. 17). No difference was apparent between WT and
MT.
3.3.2.4. Caspases 3, 6 and 7
The full forms of effector caspases 3, 6 and 7 were detected in WT and MT ovaries
throughout all investigated stages. The activation of caspase 3 and 6 remained low in
both WT and MT (Fig. 18, 19). Cleaved caspase 7 did not show a band at 20 KDa as
specified in the antibody description. We quantified instead a positive band at 16 KDa
since this band showed also in the positive control (Fig. 18). The activation of caspase 7
was low but showed a moderate increase at 19.5 dpc in both WT and MT (Fig. 20).
3.3.2.5. PARP1
The uncleaved and cleaved PARP1 were detected in WT and MT ovaries
throughout all investigated stages (Fig. 21). The ratio of the cleaved form to the full form
was high from 16.5 to 18.5 dpc (Fig. 21). At 19.5 dpc when only one ovary was used for
each lane the ratio was lower. No major differences were found between WT and MT
except for 17.5 dpc at which the cleavage of PARP1 was higher in MT, although only
one experiment was performed.
58
17.5
WT
18.5
MT
WT
19.5 dpc
MT
WT
MT
- 49 KDa
- 37 KDa
Immunoblotting of Caspase 9
(1)
(3)
(2)
(3)
Figure 14. Immunobloting (top) and activation levels (bottom ) of Caspase 9.
The number of experiments is indicated in brackets below each stage. In one experiment
four ovaries at 17.5 dpc of each genotype were used per lane.
59
17.5
WT
18.5
MT
WT
MT
21.5 dpc
WT
MT
C
NC
-35 KDa
-15 KDa
Figure 15. Immunoblotting of cleaved Caspase 9 15/35 KDa
16.5
HT
17.5
MT
HT
16.5
MT
HT
dpc
17.5
MT
HT
MT
- 51 KDa
-12-13 KDa
a
b
Figure 16. Specificity of Caspase 2L antibody
a – incubation with Caspase 2L antibody (1:1000); b – incubation with Caspase
2L 1:1000 treated in advance with blocking peptide (1:15). The anticipated full form
(51KDa) and 12-13 KDa bands decreased while other bands were intensified.
60
17.5
WT
18.5
MT
WT
19.5
MT
WT
dpc
MT
- 51 KDa
- 12-13 KDa
5112KDa
Immunoblotting of Caspase 2 L
(1)
( 3)
(2)
(3)
Figure 17. Immunobloting (top) and activation levels (bottom ) of Caspase 2L. The
number of experiments is indicated in brackets below each stage. In one experiment four
ovaries at 17.5 dpc of each genotype were used per lane.
61
17.5
WT
18.5
MT
WT
21.5 dpc
MT
WT
MT
C
NC
- 35 KDa
- 17 KDa
Immunoblotting of Caspase 3
C – control positive, NC-control negative
(1)
(2)
(1)
(2)
(1)
Figure 18. Immunobloting (top) and activation levels (bottom ) of Caspase 3. The
number of experiments is indicated in brackets below each stage.
62
17.5
WT
18.5
MT
WT
21.5 dpc
MT
WT
MT
C
NC
- 35 KDa
- 15 KDa
Immunoblotting of Caspase 6
C – control positive, NC-control negative
(1)
(2)
(1)
(2)
(1)
Figure 19. Immunobloting (top) and activation levels (bottom ) of Caspase 6. The
number of experiments is indicated in brackets below each stage.
63
17.5
WT
18.5
MT
WT
21.5 dpc
MT
WT
MT
C
NC
- 35 KDa
- 20 KDa
- 16 KDa
Immunoblotting of Caspase 7
C – control positive, NC-control negative
(1)
(1)
(2)
(1)
Figure 20. Immunobloting (top) and activation levels (bottom ) of Caspase 7. The
number of experiments is indicated in brackets below each stage.
64
16.5
WT
MT
17.5
WT
18.5
MT
WT
19.5 dpc
MT
WT
MT
C
- 116 KDa
- 89 KDa
Immunoblotting of PARP 1
C – control positive
(1)
(1)
(1)
(1)
Figure 21. Immunoblotting (top) and cleavage levels (bottom) of PARP1. The
number of experiments is indicated in brackets below each stage.
65
3.3.4. Localization of cleaved Caspase 9 (35/15 KDa and 37 KDa), cleaved
Caspase 7 (20 KDa) and cleaved PARP1 (24 KDa) in ovaries at 18.5 dpc
The results of western blot experiments showed that the average activation level of
caspase 9 was maximum at 18.5 dpc in both WT and MT. Therefore we chose this stage
to study the localization of two forms of cleaved Caspase 9 (35/15 KDa and 37 KDa) and
cleaved PARP1 (24 KDa). The specificity of antibody of 35/15 KDa form of Caspase 9
was confirmed by western-blot (Fig 15). The oocytes were identified by double
immunolabelling with the anti-GCNA1 antibody.
3.3.4.1. Localization of the 35/15 KDa cleaved forms of Caspase 9
The 35 KDa fragment of caspase 9, a product of initial caspase 9 activation, was
localized in the cytoplasm of most oocytes and in the nucleus of much fewer oocytes in
both WT and MT ovaries (Fig. 22 a-f). We also observed cleaved 35/15 KDa caspase 9
staining in the cytoplasm of cells which were negative for GCNA1 especially in MT
(Fig. 22 d-f). The staining of cleaved caspase 9 was diffuse in both cytoplasm and
nucleus.
3.3.4.2. Localization of 37 KDa of the cleaved form of Caspase 9
The 37 KDa fragment of caspase 9, a product of effector caspases, was seen
mainly in the nuclei, which were smaller than typical nuclei of oocytes and negative for
GCNA1 (Fig. 23 a-f). Only one cell with weak GCNA1 and cleaved caspase 9 labelling
was seen on one section. Therefore, the cell type of the majority of positive cells could
not be identified. These cells may represent somatic cells or germ cell, which have lost
GCNA1. Occasionally, positive caspase 9 cells (Fig. 23 g-i) but negative for GCNA1
were observed protruding the surface epithelium. Weak cytoplasmic staining was also
observed, more intense in WT.
3.3.4.3. Localization of cleaved Caspase 7
The antibody for cleaved Caspase 7 showed a weak staining in the cytoplasm of
most oocytes when used at a 1:100 or 1:300 dilution. The staining was more intense
when 1:50 dilution was used.
66
3.3.4.4. Localization of cleaved PARP1 (24 KDa)
The cleaved PARP1 was predominantly seen in the nucleus of some oocytes (Fig.
24 a-b). GCNA1 labelling was always weaker in PARP1 positive oocytes (Fig. 24 c-e).
We observed also PARP1 positive cells negative for GCNA1 (Fig. 24 f-h). PARP1
positive cells were seen more frequently in a MT ovary comparing to the WT control.
We observed oocytes protruding from the surface epithelium which were negative for the
cleaved PARP1.
67
a
b
c
d
e
f
Figure 22. Double immunolabelling of GCNA1 (green) and the 35/15 KDa cleaved forms of caspase 9 (red) in ovarian sections.
The yellow color is due to autofluorescence in blood cells. The bar represents 50 µm.
(a) Immunolabelling of GCNA1 and (b) the 35/15 KDa fragments of caspase 9 in a section of WT ovary at 18.5 dpc. (c) Merged. The
35 KDa caspase 9 is present in the cytoplasm of most oocytes (arrows) and in the nucleus of some oocytes (arrowheads)
(d) Immunolabelling of GCNA1 and (e) the 35/15 KDa fragments of caspase 9 in ovarian section of MT ovary at 18.5 dpc. (f)
Merged. The 35 KDa caspase 9 is present in the cytoplasm of oocytes (arrows) but also in cells which are negative for GCNA1
(arrowheads)
68
Figure 23. Double immunolabelling of GCNA1 (green) and the 37 KDa cleaved
form of Caspase 9 (red) in ovarian sections. The yellow color is due to
autofluorescence in blood cells. The bar represents 50 µm in a-c, 20 µm in d-f and 100
µm in g-i
(a) Immunolabelling of GCNA1 and (b) the 37 KDa fragment of caspase 9 in a section
of MT ovary at 18.5 dpc. (c) Merged. Intense labelling of 37 KDa caspase 9 is
predominantly seen in the nucleus, which is smaller than typical oocytes and negative for
GCNA1 (arrowheads). One positive 37 KDa caspase 9 cell showed a weak GCNA1
staining (arrow)
(d) High magnification of cells negative for GCNA1 but (e) positive for 37KDa caspase
(arrowheads). (f) Merged
(g) GCNA1 negative cells but (h) positive for cleaved Caspase 9 (37 KDa) (arrowhead)
near the surface epithelium. (i) Merged
69
a
b
c
d
e
f
g
h
i
70
a
b
c
d
e
f
g
h
Figure 24. Double immunolabelling of GCNA1 (green) and the 24 KDa cleaved
form of PARP1 (red) in ovarian sections. The yellow color is due to autofluorescence
in blood cells. The bar represents 20 µm in a-b and 100 µm in c-h.
(a) Immunolabelling of GCNA1 and cleaved PARP1 in sections from WT (PARP1 1:50)
and (b) MT (PARP1 1:100) ovaries
(c) High magnification of an oocyte positive for GCNA1 (arrowhead) and for (d) cleaved
PARP1 (arrowhead). (e) Merged
(f) High magnification of a cell negative for GCNA1 and (g) positive for cleaved PARP1
(arrowhead). (h) Merged
71
4. Discussion
4.1. Variation in the number of germ cells in fetal and neonatal ovaries during
meiotic prophase progression
We observed a marked loss of oocytes in MT comparing to the WT at
20.5 dpc on histological sections and by 22.5 dpc very few oocytes were seen in
MT. This observation is in accordance with Edelmann et al. (1999) who reported
that at day 3 pp ovaries of Msh5-/- females contained a few oocytes.
By
counting the total number of germ cells in cell suspension from the whole ovary
we observed that the loss of oocytes in MT was constantly larger comparing to
the control WT and HT from 16.5 to 20.5 dpc.
This may suggest that a
pachytene checkpoint monitoring the unrepaired DSBs or the persistence of
repair intermediates may be active throughout meiotic prophase eliminating
defective oocytes in MT. We found a significant difference between MT and WT
at the day of birth at 19.5 dpc. We do not exclude the possibility that germ cells
were lost during the preparation of cell suspensions; however this should not
have affected the comparison between WT/HT and MT. We observed that
meiotic defect in MT oocytes occurred at zygotene and ɣH2AX persisted in the
defective oocytes up to 19.5 dpc suggesting the persistence of unrepaired DSBs
due to the incomplete synapsis and crossing-over. The MT oocytes survived into
post-zygotene stages suggesting that there is a delay between the appearance of
meiotic defect and the elimination of oocytes. It is speculated that ɣH2AX may
be a component of the pachytene checkpoint (Burgoyne et al. 2007) and its
persistent presence in MT oocytes suggests a role in their recognition and
elimination.
The mechanism of germ cell elimination in Msh5 null mutant females in
relation to meiotic progression has been discussed by Morelli and Cohen (2005).
Their interpretation is that “the meiotic disruption in Msh4-/- and Msh5-/- mice
occurs between embryonic days 16 and 18, yet the oocyte loss is not observed
until shortly after birth. This would suggest either that earlier stages of prophase I
72
are prolonged in the absence of these MutS homologs, or that detection of failed
synapsis events is not triggered immediately, perhaps as a result of a gonad-wide
monitoring system for tracking ovarian development, or by cell- intrinsic
checkpoint systems that activate at birth”. Our results support their hypothesis
except that we found that oocytes loss occurs also before birth in MT ovaries.
4.3. Apoptotic pathway
We found a high level of activation of the initiator caspase 9 in fetal and
neonatal ovaries of both WT and MT by using an antibody which recognises the
full form 49 KDa and a cleaved form at 37 KDa. The average level was higher at
18.5 dpc when most of the oocytes in WT reached the pachytene or diplotene
stage of meiosis.
It was reported that the 37 KDa fragment of caspase 9 is a feed-back
product of the effector caspase 3 (Srinivasula et al. 1998). Our results showed
that this fragment is localized mainly in the nuclei of cells negative for GCNA1.
Therefore the cell type could not be identified. One possibility is that the cells
harboring the 37 KDa fragment of caspase 9 represent oocytes in advanced
phases of apoptosis in which GCNA1 protein has been lost. On the other hand,
we observed that the cleaved 35/15 KDa form, the product of initial caspase 9
activation (Srinivasula et al. 1998), was present in the cytoplasm of many oocytes
in WT and MT but also in some other cells which were negative for GCNA1. We
also observed positive staining in the nuclei of some oocytes. The presence of
35/15 KDa fragments in the cytoplasm of many oocytes may suggest that caspase
9 is constitutively activated and may translocate into the nucleus with
advancement in apoptosis. Our observations were made only on ovaries of 18.5
dpc. Immunolocalization at earlier and later stages would provide more
information on the involvement of both forms of caspase. The specificity of
35/15 KDa fragments were confirmed by western blot experiments. However it
would be more convincing to use a blocking peptide in immunolocalization
studies.
73
By western blot, caspase 2L was found to be activated in a similar pattern
to Caspase 9 but the level of activation was very low in both WT and MT.
Without immunolocalization we cannot conclude if caspase 2 is involved in the
elimination of oocytes during meiotic prophase.
It has been shown that caspase 9 can process procaspase 3 and 7 but not
procaspase 6 (Srinivasula et al. 1998). Our results indicate that the level of
activation of effector caspases 3 and 6 was very low in both WT and MT at all
investigated stages of development and probably they are not involved in the
execution of apoptosis in fetal and neonatal ovaries. Effector caspase 7 also
showed a low level of activation but we observed an increase at 19.5 dpc in both
WT and MT. Therefore, caspase 7 is a possible candidate as an effector caspase
in neonatal ovaries. Our first experiments on immunolabelling of ovarian sections
with cleaved caspase 7 showed a rather faint cytoplasmic staining in many cells
either positive or negative for GCNA1. However our results on both activation
and localization of caspase 7 are preliminary and additional experiments are
necessary to decide if this caspase is involved.
By western blot we found a high level of PARP1 cleavage from 16.5 to
18.5 dpc in both WT and MT. At 17.5 dpc we observed a higher level of cleavage
of PARP1 but further experiments are needed for confirmation. In ovarian
sections, cleaved PARP1 was seen exclusively in nuclei of both GCNA1 positive
and negative cells. The negative GCNA1 cells may represent oocytes which had
lost the GCNA1 protein due to advanced apoptosis.
In experiments by western blot we did not find major differences between
WT and MT. This may be due to technical limitation of using whole ovaries
instead of isolated oocytes. We observed in some cases that both the full form
and the cleaved form of caspases appeared to be higher in MT comparing to the
WT at the same stage of development. This may suggest an upregulation of full
form of caspases in MT.
It may be more informative if we could normalize the protein levels by using
germ cell marker proteins. The use of housekeeping proteins such actins would
not be helpful since the proportion of germ cells may differ between WT and MT
74
ovaries. It is, however, also conceivable that the elimination of oocytes is too
rapid to be captured by the detection of intermediate apoptotic molecules.
5. Conclusion
Our results suggest that a checkpoint which recognizes the unrepaired
DSBs operates in the elimination of MT oocytes during meiosis.
A mitochondrial apoptotic pathway involving caspase 9 and cleavage of
PARP1 may be activated in the oocytes during meiotic prophase progression in
both WT and MT ovaries. Whether caspase 7 plays a role as an effector caspase
in this pathway needs further confirmation. However, the excessive loss in MT
ovaries does not reflect in the overall levels of caspase activation. We cannot
exclude the possibility that other types of cell death, such as autophagy, act in
parallel in elimination of Msh5 -/- oocytes in prenatal and neonatal ovaries. The
involvement of caspase 7 and 9 in the loss of oocytes should be confirmed by
studies on germ cells in Caspase 7 and Caspase 9 deficient mice. It would be also
informative to study if MT oocytes are rescued in the absence of Caspase 9,
Caspase 7 or Caspase 2.
75
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