ABSTRACT In mammals, the pool of primordial follicles at birth is determinant for female fertility. Exposure to IR during oogonia proliferation and the diplotene stages of ovarian

development induced the virtual disappearance of primordial follicles in the postnatal ovary, while half the follicular reserve remained present after irradiation during the

zygotene/pachytene stages. This sensitivity difference was correlated with the level of caspase-2 expression evaluated by immunohistochemistry. At the diplotene stage, Western blot and

caspase activity analysis revealed that caspase-2 was activated 2 h after irradiation and a significant increase in the number of oocytes expressing cleaved caspase-9 and -3 occurred 6 h

SIMILAR CONTENT BEING VIEWED BY OTHERS ENHANCED PRO-APOPTOSIS GENE SIGNATURE FOLLOWING THE ACTIVATION OF TAP63Α IN OOCYTES UPON Γ IRRADIATION Article Open access 04 March 2022 EPAS1

EXPRESSION CONTRIBUTES TO MAINTENANCE OF THE PRIMORDIAL FOLLICLE POOL IN THE MOUSE OVARY Article Open access 16 April 2024 INDIVIDUAL-OOCYTE TRANSCRIPTOMIC ANALYSIS SHOWS THAT GENOTOXIC

CHEMOTHERAPY DEPLETES HUMAN PRIMORDIAL FOLLICLE RESERVE IN VIVO BY TRIGGERING PROAPOPTOTIC PATHWAYS WITHOUT GROWTH ACTIVATION Article Open access 11 January 2021 MAIN Ovarian lifespan is

tightly limited by the size of the primordial follicle stock whose formation is initiated during fetal life and is completed at or shortly after birth.1 Despite recent data emphasizing a

possible follicular renewal in the postnatal mammalian ovary involving the entry into differentiation of germline stem cells, the pool of primordial follicles remains determinant for female

fertility.2, 3 Chemotherapy and radiotherapy have radically increased long-term survival of young cancer patients, but major side effects of these treatments are ovarian failure and

infertility.4 This premature ovarian failure results from the depletion of follicular reserves, since the nearly complete destruction of primordial follicles occurs in the young adult female

mouse ovary 2 weeks after a single exposure to 0.1 Gy of ionizing radiation (IR).5 However, the molecular mechanisms involved are still mostly unknown. Radiation exposure induces cell death

_via_ apoptosis, a genetically regulated process that is a physiological way of removing cells that are not needed or are damaged. Irradiation-induced production of DNA double-strand breaks

(DSBs) has been shown to play a central role in triggering the mitochondrial apoptotic pathway.6 However, other pathways, such as generation of ceramide from cellular membranes, can

initiate apoptosis after exposure to ionizing radiation.7 Radiation-induced recruitment of the intrinsic apoptotic pathway involves the release of cytochrome _c_ from mitochondria leading to

the sequential activation of caspase-9 and -3.6 Although many upstream components can activate the mitochondrial apoptosis pathway in response to different apoptotic stimuli,8 it appears

that caspase-2, which belongs to the initiator caspase family, plays a central role in triggering DNA damage-induced apoptosis through mitochondrial activation.9 Two different mRNA species

derived from alternative splicing encode two proteins, caspase-2L, which induces cell death, and caspase-2S, a truncated protein which can antagonize cell death.10, 11 Caspase-2 is required

in cytotoxic stress-induced apoptosis before mitochondrial apoptosis12 and is activated soon after exposure to various apoptotic stimuli, including ionizing radiation.13 Moreover,

procaspase-2 is the only procaspase present constitutively in the nucleus from which it can induce the mitochondrial apoptotic pathway.14 This suggests possible direct DNA DSB-induced

caspase-2 activation. However, cytoplasmic cleaved caspase-2 can also signal directly and indirectly to mitochondria.9 Mouse fetal ovary arises under the mesonephros at 11.5 days

postconception (dpc) as an undifferentiated anlage rapidly colonized by the primordial germ cells (PGCs). These PGCs, also termed oogonia, actively proliferate before initiating the first

meiotic prophase from 13.5 dpc onwards. Most oocytes reach the zygotene stage around 14.5 dpc and shortly after enter the pachytene stage. At 18.5 dpc the first diplotene stages are observed

in the ovary and all oocytes have reached this stage at birth (19.5 dpc). The oocytes will remain blocked at this stage until their ovulation. Around birth, somatic cells intensely

proliferate to enclose the oocyte then forming the primordial follicles. The great majority of these small follicles remain quiescent while a few immediately start growing spontaneously.15

Germ cells have long been known to be very sensitive to genotoxic stress. However, this sensitivity greatly changes according to their developmental stage. For instance, meiotic pachytene

oocytes of _Caenorhabditis elegans_ (_C. elegans_) are hyper-resistant to ionizing radiation compared to diplotene/diakinesis stage oocytes and early embryonic cells.16, 17 In the current

report, we studied the effect of radiation during the course of mouse ovarian development to characterize changes in the radiosensitivity of mammalian ovary. Moreover, we attempted to

dissect some molecular mechanisms involved in the genotoxic stress-induced apoptosis of primordial follicles, the stage identified as the most sensitive. In particular, we sought to

determine which follicular compartment is most affected by ionizing radiation, and to investigate the role of caspase-2 in the depletion of the follicular reserve after exposure to

radiation. Our data demonstrate a relative radioresistance of oocytes in the course of meiotic homologous recombination repair, while irradiation during oogonia proliferation or just after

follicle formation induces virtual destruction of the primordial follicle pool. Moreover, the high radiosensitivity of primordial follicles results essentially in a fast and massive wave of

apoptosis of quiescent oocytes involving a caspase-2-dependent activation of the mitochondrial pathway. RESULTS IR AFFECTS FOLLICLE COUNT DIFFERENTLY OVER THE COURSE OF OVARIAN DEVELOPMENT

The effect of ionizing radiation on the developing ovary was analyzed by counting the number of remaining follicles in 8 dpp ovaries after exposure to a total dose of 1. 5 Gy at 12.5, 14.5,

18.5 dpc and 1 dpp. _γ_-Radiation (1.5 Gy) during oogonia proliferation (12.5 dpc) and the diplotene/diakinesis stage of meiosis prophase I (1 dpp) greatly reduced the total number of

follicles at 8 dpp compared to control (89 and 97%, respectively, _P_⩽0.05, Figure 1a). Exposure to the same dose during the zygotene (14.5 dpc) and pachytene (18.5 dpc) stages only induced

respective decreases of 56 and 52% (_P_⩽0.05) (Figure 1a). The primordial follicle pool in the 8 dpp ovary almost disappeared after exposure to 1.5 Gy on 12.5 dpc and 1 dpp (98 and 99.7%,

respectively, _P_⩽0.05), whereas 37 and 41% (_P_⩽0.05) of primordial follicles remained when irradiation was performed on 14.5 and 18.5 dpc, respectively (Figure 1b). Moreover, 1.5 Gy at 1 

dpp greatly reduced the population of primary follicles (87%, _P_⩽0.05), but had no significant effect at 12.5, 14.5 and 18.5 dpc (Figure 1b). Secondary follicles were also affected by

ionizing radiation since a significant decrease in their number was observed at all stages when compared to control (56, 37, 28 and 84% after an exposure to 1.5 Gy at 12.5, 14.5, 18.5 dpc

and 1 dpp, respectively, _P_⩽0.05, Figure 1b). On the other hand, the number of atretic follicles increased approximately threefold after exposure to 1.5 Gy (_P_⩽0.05), except at 1 dpp where

no significant effect was observed (Figure 1b). IR INDUCES APOPTOSIS OF THE OOCYTE IN PRIMORDIAL FOLLICLES We investigated the deleterious effect of ionizing radiation on 1 dpp ovary since

this stage of development is the most sensitive to irradiation, according to our first results. To investigate which follicular compartment (i.e. somatic or germ cells) was the most

sensitive to ionizing radiation, we first realized dose response experiments to determine which was the lowest dose that eradicated virtually the whole follicle population at 1 dpp (data not

shown). This threshold was achieved with a total dose of 0.5 Gy. All further experiments were performed with the 0.5 Gy dose. Analysis of semi-thin sections of control and irradiated

ovaries evidenced compaction of oocyte nuclei from primordial follicles 9 h postirradiation (Figure 2a). Electron microscopy analysis showed that the normal primordial follicles with two or

three flattened granulosa cells possessed round ooplasmic membranes, pale germinal vesicles, and one or more nucleoli (Figure 2b). On the other hand, morphological changes characteristic of

apoptosis in the oocytes of primordial follicles, such as progressive chromatin condensation and increase in lipid droplets, were clearly observed postirradiation. Analysis of 1 dpp ovary

after exposure to IR by TUNEL showed that hardly any granulosa cells were TUNEL-positive compared to oocytes (Figure 3a). Nearly all the TUNEL-positive cells were also stained for MVH, a

specific germ cell marker. Oocytes from primordial follicles were strongly affected by IR, unlike the oocytes enclosed in growing follicles, which were weakly stained for TUNEL. The

percentage of apoptotic oocytes had increased fivefold (_P_⩽0.05) by 9 h postirradiation, and peaked at 12 and 24 h (5±0.5% at 0 h _vs_ 41.7±3.4 and 39.2±2.6% at 12 and 24 h, respectively,

_P_⩽0.05, Figure 3b). A return equivalent to the basal level was observed 48 h after irradiation. IR INDUCES APOPTOSIS IN OOCYTES ISOLATED FROM 1 DPP OVARIES To confirm the great sensitivity

of female germ cells to genotoxic stress, oocytes from 1 dpp ovary, which contains mainly primordial follicles, were isolated using a micromanipulator after enzymatic dissociation and based

on their large and round shape (Figure 4a). As shown in Figure 4b, sorted cells expressed the two oocyte markers, MSY2 and c-kit, but not FOXL2, used as marker of granulosa cells,

demonstrating the high degree of purity achieved by this isolation technique. Irradiation-induced condensation of the nuclei (a characteristic of apoptosis) of isolated oocytes was

investigated using Hoechst staining (Figure 4c). Almost all control sorted oocytes maintained a high nucleo-cytolasmic ratio with a large and round-shaped nucleus, whereas 98% of irradiated

oocytes displayed condensed and small nuclei 8 h postirradiation (Figure 4d). At 16 h after irradiation, all oocytes were dead as shown by PI-incorporation, whereas very few control oocytes

were PI-positive (Figure 4e). IR INDUCES THE ACTIVATION OF CASPASE-9 AND -3 IN PRIMORDIAL FOLLICLES We investigated the expression kinetics of cleaved caspase-3 and cleaved caspase-9

post-0.5 Gy at 1 dpp to study the involvement of the intrinsic apoptotic pathway in ionizing radiation-induced follicular depletion. As observed with TUNEL analysis, oocytes from primordial

follicles were mainly stained for the cleaved forms of caspase-3 and -9 compared to granulosa cells and growing oocytes (Figure 5a and b). Oocytes expressing cleaved caspase-3 and cleaved

caspase-9 were increased in number from 6 h postirradiation (four- and threefold, respectively, _P_⩽0.05, Figure 5a and b). More than 60% of oocytes stained for the cleaved forms of

caspase-3 and -9 between 9 and 12 h after irradiation (_P_⩽0.05, Figure 5a and b). The proportion of stained oocytes started to decrease significantly at 24 h (_P_⩽0.05) and by 48 h had

reached a level equivalent to that observed at 6 h (Figure 5a and b). IR CHANGES CASPASE-2L EXPRESSION AND PRODUCTION IN THE NEWBORN OVARY RT–PCR analysis of caspase-2 expression was carried

out using a primer set for detection of caspase-2L and -2S. The two forms of caspase-2 mRNAs were detected in the control ovary, but we observed that caspase-2L mRNA predominated, and

caspase-2S mRNA was barely detectable (Figure 6a). No change was observed in the caspase-2S/-2L ratio after exposure to _γ_-radiation (Figure 6a). Quantitative RT-PCR measurement of

caspase-2 mRNA revealed that caspase-2 expression decreased from 6 h postirradiation. This was true whether caspase-2 expression was normalized according to _β_-actin (i.e. related to the

whole ovary) or to MVH (i.e. related only to oocytes) (Figure 6b). Caspase-2L production in the ovary postirradiation was specifically studied by immunohistochemistry using an antibody that

recognizes cleaved as well as procaspase-2L. In the 1 dpp ovary (0 h), caspase-2L was detected in the cytoplasm and nucleus of oocytes, and a very weak signal was detected in some somatic

cells (Figure 6c). IR induced a strong increase in staining intensity, particularly in the oocyte nuclei (Figure 6c). A significant increase in the percentage of oocytes with caspase-2

nuclear staining was observed postirradiation (19±9.8% at 0 h _vs_ 74±10.2% and 77±9.2 at 3 and 6 h, respectively, _P_⩽0.05). CASPASE-2L IS ACTIVATED EARLY IN THE 1 DPP OVARY POSTIRRADIATION

Caspase-2L activation induced by ionizing radiation (0.5 Gy) in the 1 dpp ovary was studied by measuring the cleavage of the VDVAD-AFC substrate. The specificity of the activity was

assessed by using a specific irreversible caspase-2 inhibitor (z-VDVAD-fmk). Specific caspase-2 activity was undetectable in the control ovary and increased from 2 h postirradiation (Figure

7a). Western blot analysis showed that caspase-2L precursor (48 kDa) was present in the control 1 dpp ovary. Cleavage of the procaspase-2 to the p33 subunit was observed 2 h post-IR (Figure

7b). The proform and the processing product were also detected up to 6 h after irradiation without major change in their content and ratio (data not shown). SPECIFIC CASPASE-2 INHIBITOR

Z-VDVAD-FMK PREVENTS IR-INDUCED CASPASE-9 ACTIVATION AND INCREASES OOCYTE SURVIVAL POST-IR To analyze the involvement of caspase-2L in the follicular depletion induced by ionizing radiation,

ovaries in organ culture were exposed to IR in the presence or absence of z-VDVAD-fmk. As observed _in vivo_ (Figure 5), the number of oocytes expressing cleaved caspase-9 increased

strongly 9 h postirradiation in organ culture (78.9±2.8 _vs_ 8.6±1.8% in control, _P_⩽0.05, Figure 8a). On the other hand, pretreatment with z-VDVAD-fmk (100 _μ_M) decreased by approximately

64% (_P_⩽0.05) the number of caspase-9 positive oocytes after exposure to IR (Figure 8a). Similar results were obtained when observing cleaved caspase-3 (data not shown). The number of

surviving oocytes in the presence of z-VDVAD-fmk 48 h post-IR had sharply decreased but remained nearly twice that of untreated irradiated ovaries (577±69 _vs_ 307±75, _P_⩽0.05, Figure 8b).

DMSO (vehicle) had no significant effect compared to controls (data not shown). CASPASE-2L DOES NOT COLOCALIZE WITH ΓH2AX FOCI FOLLOWING EXPOSURE TO IR DNA DSBs induced by genotoxic stress

are considered to be the most lethal damage to the cell. Analysis of _γ_H2AX expression (a marker of DSBs) on ovarian sections by immunofluorescence revealed the appearance of many _γ_H2AX

foci in the oocytes nuclei 3 h postirradiation (0.5 Gy), whereas almost no _γ_H2AX foci were detected in somatic cells (Figure 9a). No clear colocalization of caspase-2L and _γ_H2AX foci was

observed in the germinal vesicle 3 h after exposure to IR (Figure 9b). CASPASE-2L EXPRESSION CORRELATES WITH OOCYTE RADIOSENSITIVITY THROUGHOUT OVARIAN DEVELOPMENT Caspase-2 expression was

investigated in the ovary from 12.5 dpc to 2 dpp (Figure 10).Staining of caspase-2 was faint on 18.5 dpc, intensified on 0 dpp, and was maintained thereafter. Only diplotene stage oocytes

from small quiescent follicles displayed a strong signal, while oogonia (12.5 dpc), zygotene (14.5 dpc) and pachytene (17.5) stage and diplotene stage oocytes from growing follicles (2 dpp)

bore no detectable staining. DISCUSSION This report demonstrates that the sensitivity of the mammalian ovary to IR changes throughout development, and provides new data on follicular

depletion and on apoptotic pathways activated in the oocyte after exposure to genotoxic stress. Exposure to IR during oogonia proliferation (12.5 dpc) and the diplotene/diakinesis stage of

meiosis prophase I (1 dpp) induces an almost total disappearance of the follicular reserve. This confirms previous data obtained in the rat.18, 19 Interestingly, we show that the deleterious

effect of _γ_-radiation is less marked when irradiation is performed during the zygotene/pachytene stage (14.5–18.5 dpc) in the mammalian ovary, since approximately half of the primordial

follicles remain in the ovary at 8 dpp. This low oocyte radiosensitivity at this stage could be due to a low basal ratio of pro- and antiapoptotic regulators, as described in different cell

lineages.20 Here, we demonstrate that caspase-2 expression is initiated when oocytes reach the diplotene stage. Therefore, the radiosensitivity of mouse oocytes in the course of ovarian

development appears to be closely related to the pattern of caspase-2 expression. Another explanation of radioresistance during the zygotene/pachytene stage could be the high level of

expression of the DNA repair machinery necessary for meiotic homologous recombination, as already described in _C. elegans_.16 Interestingly, oogonia radiosensitivity does not seem to be

related to caspase-2 expression, indicating that IR probably triggers the death of these mitotic cells through a different mechanism. Our results demonstrate that IR depletes the follicular

reserve in mouse newborn ovary. Our three-pronged approach (i.e. electron microscopy, TUNEL and oocyte isolation) shows that this destruction of the primordial follicle pool results

specifically from a rapid and massive wave of apoptosis of the germinal compartment. Growing oocytes were little affected compared to quiescent oocytes, in agreement with previous data

obtained in the rat ovary.19 The radiosensitivity of quiescent oocytes and the relative resistance of growing oocytes remain unexplained. As discussed above, it may be that growing and

quiescent oocytes have different basal patterns of expression of pro- and antiapoptotic regulators. Our results tend to support this idea, since caspase-2 expression was very low in growing

oocytes but strong in quiescent oocytes. One should not, however, exclude an antiapoptotic effect of the follicular environment of growing oocytes, since granulosa cells begin to produce

growth factors essential for follicle growth.21 We observed a rapid and massive activation of caspase-9 and -3 after irradiation in the female germ cells, demonstrating that the

mitochondrial pathway of apoptosis is activated in oocytes in response to genotoxic stress, as in most DNA damage-induced apoptosis models.20 Moreover, by removing female germ cells from the

ovarian environment, we show that somatic follicular cells seem to be not required for irradiation-induced apoptosis of the primordial oocytes. So, the intrinsic apoptotic pathway appears

to provide an efficient mechanism for the elimination of quiescent oocytes carrying DNA lesions. One important finding of this report is that caspase-2 is involved in genotoxic

stress-induced oocyte apoptosis. In addition to the predominant caspase-2L mRNA expression previously reported in the mouse ovary,22, 23 we show here by immunohistochemistry that this

proapoptotic factor displays oocyte-specific expression in the postnatal ovary and that exposure to genotoxic stress leads to a decrease in caspase-2L mRNA expression in the ovary. As

transcriptional activity is repressed in quiescent oocytes from primordial follicle,24 this depletion in caspase-2 mRNA could be related to the recruitment of this transcript to the

translation process. Indeed, a similar depletion of caspase-2 and other apoptotic mRNAs in response to genotoxic stress has previously been described in the ovulated oocyte known to be in a

transcriptional repression state.25 Moreover, our results demonstrate early activation of caspase-2 which precedes the activation of caspase-9 and -3, and pretreatment of ovaries with a

selective caspase-2 inhibitor (z-VDVAD-fmk) prevents the cleavage of caspase-9 and -3 in primordial oocytes and ultimately rescues some oocytes. Taken together, our data demonstrate that

caspase-2 constitutes an early step in triggering the mitochondrial apoptotic pathway in irradiated quiescent oocytes. Caspase-2 is also involved in the natural apoptotic wave occurring

during follicle formation.23 Thus, caspase-2 appears to trigger oocyte apoptosis in response to different stimuli such as genotoxic stress and cytokine insufficiency.26 Caspase-2 inhibitor

prevented the death of some oocytes at 48 h postirradiation when compared to untreated irradiated ovaries. This indicates that caspase-2 activated pathway is functional and at least

partially required for the IR-induced oocyte death. However, although activation of downstream caspase-9 and -3 was well prevented by caspase-2 inhibitor, not all of the oocytes were rescued

and most of them died when compared to control ovaries (not irradiated). This suggests that the caspase-2-activated mitochondrial pathway is not the only mechanism triggering oocyte

apoptosis after exposure to potent genotoxic stress, or that activation of an alternative pathway occurs to compensate for the loss of caspase-2 functions. In the same line, Takai _et al_27

demonstrated that primordial follicles were not protected from 4-vinylcyclohexene diepoxide (VCD)-induced death in caspase-2- or -3-deficient mice. Caspase-2 can mediate the recruitment of

the mitochondrial apoptotic pathway using different mechanisms.9 Active caspase-2 can act directly on mitochondria or cleave cytosolic Bid protein to release cytochrome _c_ and

Smac/DIABLO.28 Caspase-2 can also trigger cytochrome _c_ release and apoptosis from the nucleus through possible Bid processing.14 We observed that _γ_-radiation induced an increase in

caspase-2 immunostaining in the oocyte cytoplasm as well as in the nucleus, suggesting that caspase-2 acts both directly and indirectly on mitochondria in triggering oocyte apoptosis in

response to genotoxic stress. Caspase-2 can be recruited and activated by different mechanisms. Our results tend to demonstrate that caspase-2 activation in irradiated oocytes seems not to

be related to a direct interaction of caspase-2 with DNA DSBs revealed by _γ_H2AX foci. Interestingly, the increase in staining of caspase-2L and the depletion in caspase-2 mRNA post-IR

suggest the occurrence of a _de novo_ caspase-2 protein synthesis, which may be functionally important for its activation. It has been documented that overexpression induces dimerization of

caspase-2, which constitutes its initial activation.29, 30 This dimerization promotes the autocleavage between the large and small subunits which is necessary to trigger cell death.29, 30

However, other mechanisms could be involved in caspase-2 activation during oocyte apoptosis. Ceramide induces the mitochondrial apoptotic pathway via activation of caspase-2,31 and the

sphingomyelin pathway is a major signaling pathway that triggers the apoptosis of irradiated oocytes.5 Furthermore, a p53-dependent caspase-2 activation involving the formation of the

PIDDosome cannot be excluded in irradiated oocytes.32 Lastly, the generation of reactive oxygen species (ROS), which occurs after irradiation, has recently been demonstrated to lead to the

activation of caspase-2.33 In conclusion, we report here essential data concerning the mechanisms involved in the IR-induced depletion of primordial follicle stock occurring after birth,

which may serve to maintain genomic integrity. We demonstrate that following genotoxic stress, the disappearance of the follicular reserve involves an early caspase-2-dependent activation of

the apoptotic mitochondrial pathway in quiescent oocytes. Moreover, caspase-2 expression is well correlated with oocyte radiosensitivity throughout ovarian development, emphasizing that

this caspase may play an important role in triggering the genotoxic stress-induced apoptosis of female germ cells. MATERIALS AND METHODS ANIMALS, WHOLE-BODY _Γ_-IRRADIATION AND TISSUE

PROCESSING Female NMRI mice were housed individually under controlled photoperiod conditions (lights on 0800–2000 h) and were supplied with commercial feed and tap water _ad libitum_. The

day after an overnight mating was counted as day 0.5 postconception (0.5 dpc). Natural birth occurred on 19.5 dpc, which was counted as day 0 postpartum (0 dpp). All animal studies were

conducted in accordance with the _NIH Guide for Care and Use of Laboratory Animals_. To evaluate the effect of IR during ovarian development, animals at 12.5, 14.5, 18.5 dpc and 1 dpp were

whole-body exposed to _γ_-irradiation using a 137Cs isotopic source with a total dose of 1.5 Gy at a dose rate of 0.62 Gy/min. Fetuses were exposed to IR _in utero_ and left with their

mother after birth. Ovaries were collected at 8 dpp, fixed immediately in Bouin's fixative or in 4% paraformaldehyde for at least 1 h, dehydrated in alcohol, paraffin-embedded and then

cut into 5 _μ_m sections. The IR-induced follicular death was studied in neonate ovaries. Animals 1 dpp old were whole-body _γ_-irradiated with a total dose of 0.5 Gy at a dose rate of 0.62 

Gy/min and ovaries were collected at 0, 1, 2, 3, 4, 5, 6, 9, 12, 24 and 48 h postirradiation. For immunohistochemistry, they were processed as described above. For Western blotting, ovaries

were frozen in liquid nitrogen, then stored at −80°C until protein extraction, and for RT-PCR analysis they were frozen in RLT buffer from the RNeasy Plus Mini Kit (Qiagen, S.A.,

Courtaboeuf, France) until RNA extraction as suggested by the manufacturer. ORGAN CULTURE Ovaries were collected at 0 dpp, cut into two pieces and placed on a Millicell filter (Millipore,

Billerica, MA; pore size: 0.45 _μ_m). The filter was floated on DMEM:HAM-F12 (50% v/v) (Invitrogen, Cergy Pontoise, France) supplemented with 15 mM HEPES (Invitrogen), 40 _μ_g/ml gentamycin

(Sigma-Aldrich, Lyon, France), 1% fetal calf serum (FCS) (Invitrogen) on 24-well culture plate (Nunc), and incubated at 37°C in an atmosphere of 5% CO2. After 24 h, organs were cultured in

the presence or absence of the caspase-2 inhibitor z-VDVAD-fmk (100 _μ_M) (Santa Cruz Biotechnologies, CA, USA) for 1 h, then exposed to 0.5 Gy of _γ_-irradiation. Culture medium was

replaced with fresh medium with or without the caspase inhibitors. After 9 h or 48 h, ovaries were fixed in Bouin's fixative for at least 1 h, dehydrated in alcohol and

paraffin-embedded for immunohistochemistry analysis. CULTURE OF DISPERSED OVARIAN CELLS Ovaries were collected at 0 dpp, and then digested using 0.5 mg/ml collagenase, 20 _μ_g/ml DNase I and

1 × trypsin/EDTA (Sigma). This procedure results in detachment of the layer of granulosa cells from the oocytes, thereby enabling isolation of ovarian cells. The suspension of ovarian cells

was cultured on four-well Labtek (Nunc) in DMEM:HAM-F12 supplemented with 15 mM HEPES, 40 _μ_g/ml gentamycin, 1% FCS for 24 h to allow cell attachment and then _γ_-irradiated with a total

dose of 0.5 Gy. After replacement of culture medium, cells were fixed at 3 h postirradiation with −20°C methanol for 5 min, and then analyzed by immunofluorescence. ISOLATION OF NEONATAL

OOCYTES Oocytes from dispersed 1 dpp ovarian cells were sorted according to their round shape and larger size using a micromanipulator system (CellTram Oil, Eppendorf, Le Pecq, France)

placed on an inverted microscope (Axiovert 200, Zeiss). The purity of the germ cell fraction was assayed by RT-PCR using two specific oocyte markers: MSY2 (primer forward:

5′-CAGCCTATAGCCGCAGAGAC-3′ and primer reverse: 5′-GGTGATGCCTCGGAACAATA-3′) and c-kit (primer forward: 5′-AATGGCCTCACGAGTTCTAT-3′ and primer reverse: 5′-ATGGAGTTCACGGATGTAGA-3′) and FOXL-2

(primer forward: 5′-AAGCCCCCGTACTCGTACGTGGCGCTCATC-3′ and primer reverse: 5′-GTAGTTGCCCTTCTCGAACATGTC-3′) as somatic cell marker. HOECHST STAINING OF ISOLATED OOCYTES The number of apoptotic

sorted oocytes was measured by assessing the percentage of oocytes displaying condensed nuclei. Briefly, after isolation, oocytes were placed in DMEM:HAM-F12 supplemented with 15 mM HEPES,

40 _μ_g/ml gentamycin, 1% FCS and then _γ_-irradiated with a total dose of 0.5 Gy. After 8 h, sorted oocytes were fixed in 4% paraformaldehyde for 20 min at 4°C, washed with PBS, transferred

to Superfrost-Plus slides and air-dried. Then, oocytes were permeabilized with 0.1% Triton X-100, stained for MVH by immunofluorescence and the nuclear chromatin was counterstained with

Hoechst 33342 (5 _μ_g/ml) for 20 min. PROPIDIUM IODIDE INCORPORATION OF ISOLATED OOCYTES Sorted oocytes were placed in DMEM:HAM-F12 supplemented with 15 mM HEPES, 40 _μ_g/ml gentamycin, 1%

FCS and then _γ_-irradiated with a total dose of 0.5 Gy. After 16 h of culture, oocytes were incubated for 20 min with propidium iodide (PI) (1 _μ_g/ml) and Hoechst 33342 (1 _μ_g/ml) to

assess the percentage of dead cells after irradiation. QUANTIFICATION OF FOLLICLE NUMBER Five _μ_m sections were mounted on glass slides and stained with hematoxylin–eosin. The follicles

were counted in every tenth section using the oocyte nucleus as a marker, and the stage of follicular development was determined as previously described.18 Briefly, follicles were classified

according to the shape and number of layers of somatic cells that surrounded the oocyte: primordial with flattened cells, primary with one layer of cuboidal cells and secondary with two

partial or complete layers of cells. Atretic follicles were identified due to the presence of a degenerating oocyte or of several pyknotic cells. ELECTRONIC MICROSCOPY Ovaries were fixed in

2.5% glutaraldehyde in cacodylate buffer before being postfixed in 1% OsO4. They were then dehydrated in graded alcohols followed by propylene oxide. Embedding was carried out in Durkupan

(Fluka). Semi-thin sections were stained with toluidine blue. Ultrathin sections were counterstained with uranyl acetate and lead citrate. Observations were made using a Jeol–1010 electron

microscope. IMMUNOHISTOCHEMISTRY Tissue sections (5 _μ_m) were mounted on glass slides and boiled for 10 min in 10 mM Tris pH 10.6 for antigen unmasking. Endogenous peroxidase activity was

blocked with 3% hydrogen peroxide for 10 min. Slides were incubated with the primary antibody overnight at 4°C. The primary antibodies used were rabbit anticleaved caspase-3 Asp 175 (diluted

1:100) (Cell Signaling Technology, Beverly, MA, USA), rabbit anticleaved caspase-9 Asp 353 (diluted 1:100) (Cell Signaling Technology), and rabbit anticaspase-2L sc-626 (diluted 1:250)

(Santa Cruz Biotechnologies). After washing in PBS, slides were incubated for 30 min at room temperature with a biotinylated goat anti-rabbit antibody diluted 1:200 (Vector Laboratories,

Peterborough, England), and then for 30 min with the avidin–biotin–peroxidase complex (Vector Laboratories). Peroxidase activity was visualized using 3,3′-diaminobenzidine (DAB) as

substrate. Finally, slides were counterstained with hematoxylin. CASPASE-2 ACTIVITY ASSAY Caspase-2 activity in ovarian lysates following exposure to IR was assayed using the Fluorometric

Caspase-2 Assay Kit (Calbiochem, La Jolla, CA, USA) according to the manufacturer's instructions. Briefly, ovarian lysates were incubated for 2 h at 37°C with VDVAD-AFC substrate (50 

_μ_M) in the presence or absence of z-VDVAD-fmk (100 nM) in the reaction buffer. Cleavage of the fluorogenic peptide substrate was monitored by AFC release using 400 nm excitation and 505 nm

emission wavelengths. Specific caspase-2 activity was considered to be the VAVADase. The total protein concentration was determined for each sample and data were normalized to the quantity

of total protein used for the assay. TERMINAL DEOXYNUCLEOTIDYLTRANSFERASE-MEDIATED DEOXYURIDINE 5′-TRIPHOSPHATE-FLUORESCEIN NICK END LABELING Apoptotic cells were detected on 4%

paraformaldehyde-fixed ovarian sections (5 _μ_m) using the _in situ_ Cell Death Detection Kit, POD (Roche, Diagnostics, Switzerland) according to the manufacturer's instructions.

IMMUNOFLUORESCENCE Fixed ovarian cells or fixed ovarian sections were incubated for 1 h with the primary antibodies at room temperature. The primary antibodies used were mouse anti-_γ_H2AX

(diluted 1:200) (Upstate – Cell Signaling Solutions), rabbit anti-MVH (diluted 1:500) (Abcam) and rabbit anticaspase-2L sc-626. After washing in PBS, cells were incubated for 45 min with a

donkey anti-mouse IgG-FITC (diluted 1:200) (Jackson ImmunoResearch Laboratories) or with a donkey anti-rabbit-Cy3 (diluted 1:1000) (Jackson ImmunoResearch Laboratories). Slides were mounted

with Vectashield with or without Dapi (Vector Laboratories) and analyzed by conventional immunofluorescence microscopy using a Provis AX70 Olympus microscope. RNA EXTRACTION AND RT–PCR Total

RNA from 1 dpp ovaries was extracted using the RNeasy Plus Mini Kit (Qiagen), and 500 ng of total RNA were reverse-transcribed using the Omniscript Reverse Transcription kit (Qiagen)

according to the kit instructions. PCR was performed to study the expression of caspase-2 isoforms after _γ_-irradiation. Primers for caspase-2L and -2S amplification were: forward

5′-TTTTTCGACTTTTTGACAATGC-3′ and reverse 5′-GCATGTCACAAGCTCTTTCAG-3′; and for _β_-actin amplification, used as reference: forward 5′-AAGAGAGGTATCCTGACCCTG-3′ and reverse

5′-GGCCATCTCCTGCTCGAAGT-3′. PCR was performed using 1 _μ_l cDNA, 3 pmol of each primer (Invitrogen), 0.2 mM of each dNTP (Invitrogen), 2 mM MgCl2 (Invitrogen), 1 U Taq DNA polymerase

(Invitrogen) in the reaction buffer (20 mM Tris-HCl (pH 8.4), 50 mM KCl) in a final volume of 25 _μ_l. PCR conditions were: (3 min/94°C), 30 cycles (45 s/94°C, 1 min/56°C, 1 min/72°C) for

caspase-2 and (2 min/94°C), 26 cycles (45 s/94°C, 50 s/50°C, 1 min/72°C) for _β_-actin. Amplification products were separated on 2 or 3% agarose gel stained with ethidium bromide (Sigma) (1 

_μ_g/ml). REAL-TIME QUANTITATIVE PCR Real-time PCRs were performed using the ABI Prism 7000 Sequence Detection system (Applied Biosystems, Courtaboeuf, France) according to the

manufacturer's instructions. Primers for caspase-2 quantification were: forward 5′-CCACAGATGCTACGGAACA-3′ and reverse 5′-GCTGGTAGTGTGCCTGGTAA-3′; for _β_-actin, used as reference:

forward 5′-TGACCCAGATCATGTTTGAGA-3′ and reverse 5′-TACGACCAGAGGCATACAGG-3′; and for MVH, used also as reference: forward 5′GAAGAAATCCAGAGGTTGGC-3′ and reverse 5′GAAGGATCGTCTGCTGAACA-3′.

Caspase-2, _β_-actin and MVH mRNAs were quantified using the SYBR Green Universal PCR Master Mix 2 × (Applied Biosystem) in a total volume of 30 _μ_l. Samples were heated for 10 min at 95°C,

followed by 40 cycles of 15 s at 95°C then 1 min at 60°C. The statistical significance of differences in mRNA expression was analyzed by the Relative Expression Software Tool (REST).34 PCR

efficiencies for caspase-2, _β_-actin and MVH were 2.07, 1.98 and 1.89, respectively. PROTEIN EXTRACTION AND WESTERN BLOTTING Ovaries (1 dpp) were resuspended in homogenization buffer (20 mM

Tris base pH 8.0 (Sigma) containing 150 mM NaCl (Sigma), 0.5 mM EDTA (Sigma), 1% Triton X 100 (Sigma), 0.1% sodium dodecyl sulfate (SDS) (Sigma), 10 mM sodium fluoride (NaF) (Sigma), 1 mM

sodium orthovanadate (NaO) (Sigma), 10 mM _β_-glycerophosphate (Sigma) and protease inhibitor cocktail (1 tablet/10 ml extraction solution, (Roche)). Samples were placed on ice for 30 min,

homogenized every 5 min and then centrifuged at 4°C for 15 min at 11 000 g to remove cellular debris, and the protein concentration in the supernatant was measured by the Bradford method.

Finally, samples were frozen at −20°C until Western blotting. Total proteins (50 _μ_g) were separated on a 12% polyacrylamide denaturing gel using Tris/glycine/SDS running buffer (25 mM Tris

base (Sigma), 200 mM glycine (pH 8.3) (Sigma), 0.1% SDS (Sigma)) and were blotted onto PVDF membranes (Amersham Biosciences, Saclay, France). Membranes were blocked for 1 h at room

temperature in Tris-buffered saline (TBS) pH 7.4 containing 0.05% Tween 20 (Sigma) and 5% non-fat dried milk, then incubated overnight at 4°C with a mouse anticaspase-2 (Cell Signaling

Technologies) diluted 1:100 in the blocking solution. Membranes were incubated for 1 h at room temperature with a goat anti-mouse IgG:HRP-linked antibody (Amersham) diluted 1:5,000 in the

blocking solution. Antibody–protein complexes were visualized using the enhanced chemiluminescence (ECL) visualization system (Amersham). Membranes were stripped to detect actin used as

reference. Briefly, they were incubated for 15 min in a stripping buffer (62.5 mM Tris-HCl pH 6.8 (Sigma), 2% SDS (Sigma), 100 mM _β_-mercaptoethanol (Sigma)). Actin expression was detected

as described above using a mouse monoclonal antiactin (CP01) antibody (Calbiochem). DATA ANALYSIS Each data point represents the mean±S.E.M. of at least three independent experiments. Images

show a representative experiment that was repeated at least three times. Data were analyzed using Graphpad Instat 3.0, by one-way ANOVA followed by the Tukey-Kramer multiple comparisons

test. ABBREVIATIONS * DAB, 3: 3′-diaminobenzidine * dpc: days postconception * dpp: days postpartum * DSBs: double-strand breaks * IR: ionizing radiation * MVH: mouse vasa homolog * PI:

propidium iodide * z-VDVAD-fmk: Z-Val-Asp(O-Me)-Val-Ala-Asp(O-Me) fluoromethyl ketone REFERENCES * Skinner MK . Regulation of primordial follicle assembly and development. _Hum Reprod

Update_ 2005; 11 (5): 461–471. Article  PubMed  Google Scholar  * Johnson J, Canning J, Kaneko T, Pru JK, Tilly JL . Germline stem cells and follicular renewal in the postnatal mammalian

ovary. _Nature_ 2004; 428: 145–150. Article  CAS  PubMed  Google Scholar  * Johnson JL, Bagley J, Skaznik-Wikiel M, Lee HJ, Adams GB, Niikura Y _et al_. Oocyte regeneration in adult

mammalian ovaries by putative germ cells in bone marrow and peripheral blood. _Cell_ 2005; 122: 303–315. Article  CAS  PubMed  Google Scholar  * Meirow D, Nugent D . The effects of

radiotherapy and chemotherapy on female reproduction. _Hum Reprod Update_ 2001; 7 (6): 535–543. Article  CAS  PubMed  Google Scholar  * Morita Y, Perez GI, Paris F, Miranda SR, Ehleiter D,

Haimovitz-Friedman A _et al_. Oocyte apoptosis is suppressed by disruption of the acid sphingomyelinase gene or by sphingosine-1-phosphate therapy. _Nat Med_ 2000; 6 (10): 1109–1114. Article

  CAS  PubMed  Google Scholar  * Prise KM, Schettino G, Folkard M, Held KD . New insights on cell death from radiation exposure. _Lancet Oncol_ 2005; 6: 520–528. Article  CAS  PubMed  Google

Scholar  * Haimovitz-Friedman A, Kan CC, Ehleiter D, Persaud RS, McLoughlin M, Fuks Z _et al_. Ionizing radiation acts on cellular membranes to generate ceramide and initiate apoptosis. _J

Exp Med_ 1994; 180: 525–535. Article  CAS  PubMed  Google Scholar  * Wang X . The expanding role of mitochondria in apoptosis. _Genes Dev_ 2001; 15: 2922–2933. CAS  PubMed  Google Scholar  *

Zhivotovsky B, Orennius S . Caspase-2 function in response to DNA damage. _Biochem Biophys Res Commun_ 2005; 331: 859–867. Article  CAS  PubMed  Google Scholar  * Wang L, Miura M, Bergeron

L, Zhu H, Yuan J . Ich-1, an Ice/ced-3-related gene, encodes both positive and negative regulators of programmed cell death. _Cell_ 1994; 78: 739–750. Article  CAS  PubMed  Google Scholar  *

Kumar S, Kinoshita M, Dorstyn L, Noda M . Origin, expression and possible functions of the two alternatively spliced forms of the mouse Nedd2 mRNA. _Cell Death Differ_ 1997; 4: 378–387.

Article  CAS  Google Scholar  * Lassus P, Opitz-Araya X, Lazebnik Y . Requirement for caspase-2 in stress-induced apoptosis before mitochondrial permeabilization. _Science_ 2002; 297:

1352–1354. Article  CAS  PubMed  Google Scholar  * Harvey NL, Butt AJ, Kumar S . Functional activation of Nedd2/ICH-1 (caspase-2) is an early process in apoptosis. _J Biol Chem_ 1997; 272

(20): 13134–13139. Article  CAS  PubMed  Google Scholar  * Paroni G, Henderson C, Schneider C, Brancolini C . Caspase-2 can trigger cytochrome c release and apoptosis from the nucleus. _J

Biol Chem_ 2002; 277 (17): 15147–15161. Article  CAS  PubMed  Google Scholar  * McClellan KA, Gosden R, Taketo T . Continuous loss of oocytes throughout meiotic prophase in the normal mouse

ovary. _Dev Biol_ 2003; 258: 334–348. Article  CAS  PubMed  Google Scholar  * Takanami T, Mori A, Takahashi H, Higashitani A . Hyper-resistance of meiotic cells to radiation due to a strong

expression of a single recA-like gene in _Caenorhabditis elegans_. _Nucleic Acids Res_ 2000; 28 (21): 4232–4236. Article  CAS  PubMed  PubMed Central  Google Scholar  * Rinaldo C,

Bazzicalupo P, Ederle S, Hilliard M, La Volpe A . Roles for _Caenorhabditis elegans_ rad-51 in meiosis and in resistance to ionizing radiation during development. _Genetics_ 2002; 160:

471–479. CAS  PubMed  PubMed Central  Google Scholar  * Mazaud S, Guigon CJ, Lozach A, Coudouel N, Forest MG, Coffigny H _et al_. Establishment of the reproductive function and transient

fertility of female rats lacking primordial follicle stock after fetal-irradiation. _Endocrinology_ 2002; 143: 4775–4787. Article  CAS  PubMed  Google Scholar  * Guigon CJ, Mazaud S, Forest

MG, Brailly-Tabard S, Coudouel N, Magre S . Unaltered development of the initial follicular waves and normal pubertal onset in female rats after neonatal deletion of the follicular reserve.

_Endocrinology_ 2003; 144: 3651–3662. Article  CAS  PubMed  Google Scholar  * Norbury CJ, Zhivotovsky B . DNA damage-induced apoptosis. _Oncogene_ 2004; 23: 2797–2808. Article  CAS  PubMed 

Google Scholar  * Van Den Hurk R, Zhao J . Formation of mammalian oocytes and their growth, differentiation and maturation within ovarian follicles. _Theriogenology_ 2005; 63 (6): 1717–1751.

Article  CAS  PubMed  Google Scholar  * Flaws JA, Kugu K, Trbovich AM, DeSanti A, Tilly KI, Hirshfield AN _et al_. Interleukin-a beta-converting enzyme-related proteases (IRPs) and

mammalian cell death: dissociation of IRP-induced oligonucleosomal endonuclease activity from morphological apoptosis in granulosa cells of the ovarian follicle. _Endocrinology_ 1995; 136:

5042–5053. Article  CAS  PubMed  Google Scholar  * Bergeron L, Perez GI, Macdonald G, Shi L, Sun Y, Jurisicova A _et al_. Defects in regulation of apoptosis in capase-2-deficient mice.

_Genes Dev_ 1998; 12: 1304–1314. Article  CAS  PubMed  PubMed Central  Google Scholar  * Serafica MD, Goto T, Trounson AO . Transcripts from a human primordial follicle cDNA library. _Hum

Reprod_ 2005; 20 (8): 2074–2091. Article  CAS  PubMed  Google Scholar  * Jurisicova A, Lee HJ, D'Estaing SG, Tilly J, Perez GI . Molecular requirements for doxorubicin-mediated death in

murine oocytes. _Cell Death Differ_ 2006; 13: 1466–1474. Article  CAS  PubMed  Google Scholar  * Morita Y, Maravei DV, Bergeron L, Wang S, Perez GI, Tsutsumi O _et al_. Caspase-2 deficiency

prevents programmed germ cells death resulting from cytokine insufficiency but not meiotic defects caused by loss of ataxia telangiectasia-mutated (Atm) gene function. _Cell Death Differ_

2001; 8: 614–620. Article  CAS  PubMed  Google Scholar  * Takai Y, Canning J, Perez GI, Pru JK, Schlezinger JJ, Sherr DH _et al_. Bax, caspase-2, and caspase-3 are required for ovarian

follicle loss caused by 4-vinylcyclohexene diepoxide exposure of female mice _in vivo_. _Endocrinology_ 2003; 144: 69–74. Article  CAS  PubMed  Google Scholar  * Guo Y, Srinivasula SM,

Druilhe A, Fernandes-Alnemri T, Alnemri ES . Caspase-2 induces apoptosis by releasing proapoptotic proteins from mitochondria. _J Biol Chem_ 2002; 277 (16): 13430–13437. Article  CAS  PubMed

  Google Scholar  * Butt AJ, Harvey NL, Parasivam G, Kumar S . Dimerization and autoprocessing of the Nedd2 (caspase-2) precursor requires both the prodomain and the carboxyl-terminal

regions. _J Biol Chem_ 1997; 273 (12): 6763–6768. Article  Google Scholar  * Baliga BC, Read SH, Kumar S . The biochemical mechanism of caspase-2 activation. _Cell Death Differ_ 2004; 11:

1234–1241. Article  CAS  PubMed  Google Scholar  * Lin CF, Chen CL, Chang WT, Jan MS, Hsu LJ, Wu RH _et al_. Sequential caspase-2 and caspase-8 activation upstream of mitochondria during

ceramide- and etoposide-induced apoptosis. _J Biol Chem_ 2004; 279 (39): 40755–40761. Article  CAS  PubMed  Google Scholar  * Tinel A, Tschopp J . The PIDDosome, a protein complex implicated

in activation of caspase-2 in response to genotoxic stress. _Science_ 2004; 304: 843–846. Article  CAS  PubMed  Google Scholar  * Prasad V, Chandele A, Jagtap JC, Kumar PS, Shastry P .

ROS-triggered caspase 2 activation and feedback amplification loop in beta-carotene-induced apoptosis. _Free Radic Biol Med_ 2006; 41 (3): 431–442. Article  CAS  PubMed  Google Scholar  *

Pfaffl MW, Horgan GW, Dempfle L . Relative expression software tool (REST©) for groupe-wise comparison and statistical analysis of relative expression results in real-time PCR. _Nucleic

Acids Res_ 2002; 30: e36. Article  PubMed  PubMed Central  Google Scholar  Download references ACKNOWLEDGEMENTS This work was supported by Electricité De France (EDF). We thank V Neuville, S

Leblay and C Chauveau for taking care of the animals, A Gouret for her secretarial help and D Marsh for his useful critical review of this manuscript. AUTHOR INFORMATION AUTHORS AND

AFFILIATIONS * CEA, DSV/DRR/SEGG/LDRG, Laboratory of Differentiation and Radiobiology of the Gonads, Unit of Gametogenesis and Genotoxicity, F-92265 Fontenay aux Roses, France V Hanoux, C

Pairault, R Habert & G Livera * Univ Paris 7 – Denis Diderot, U.F.R of Biology, UMR-S 566, Fontenay aux Roses, F-92265, France V Hanoux, C Pairault, R Habert & G Livera * INSERM,

U566, Fontenay aux Roses, F-92265, France V Hanoux, C Pairault, R Habert & G Livera * Institute of Experimental Morphology & Anthropology, Bulgarian Academy of Sciences, Sofia,

Bulgaria M Bakalska Authors * V Hanoux View author publications You can also search for this author inPubMed Google Scholar * C Pairault View author publications You can also search for this

author inPubMed Google Scholar * M Bakalska View author publications You can also search for this author inPubMed Google Scholar * R Habert View author publications You can also search for

this author inPubMed Google Scholar * G Livera View author publications You can also search for this author inPubMed Google Scholar CORRESPONDING AUTHOR Correspondence to G Livera.

ADDITIONAL INFORMATION Edited by JL Tilly RIGHTS AND PERMISSIONS Reprints and permissions ABOUT THIS ARTICLE CITE THIS ARTICLE Hanoux, V., Pairault, C., Bakalska, M. _et al._ Caspase-2

involvement during ionizing radiation-induced oocyte death in the mouse ovary. _Cell Death Differ_ 14, 671–681 (2007). https://doi.org/10.1038/sj.cdd.4402052 Download citation * Received: 05

April 2006 * Revised: 04 September 2006 * Accepted: 18 September 2006 * Published: 03 November 2006 * Issue Date: 01 April 2007 * DOI: https://doi.org/10.1038/sj.cdd.4402052 SHARE THIS

ARTICLE Anyone you share the following link with will be able to read this content: Get shareable link Sorry, a shareable link is not currently available for this article. Copy to clipboard

Provided by the Springer Nature SharedIt content-sharing initiative KEYWORDS * caspase-2 * oocyte * ionizing radiation * genotoxic stress * apoptosis * mitochondrial pathway