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BOR - Papers in Press, published online ahead of print July 30, 2004.
Biol Reprod 2004, 10.1095/biolreprod.104.033589
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BIOLOGY OF REPRODUCTION 71, 1936–1942 (2004)
DOI: 10.1095/biolreprod.104.033589
© 2004 by the Society for the Study of Reproduction, Inc.

Gamete Biology

Mitochondrial Dysfunction in Mouse Oocytes Results in Preimplantation Embryo Arrest in Vitro1

George A. Thouas2,3,4, Alan O. Trounson4, Ernst J. Wolvetang3, and Gayle M. Jones3,4

Monash Institute of Reproduction and Development,3 Monash University, Clayton, Victoria 3168, Australia Monash Immunology and Stem Cell Laboratories,4 Monash University, Clayton, Victoria 3800, Australia


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Oocyte mitochondrial dysfunction has been proposed as a cause of high levels of developmental retardation and arrest that occur in human preimplantation embryos generated using assisted reproductive technology in the treatment of some causes of female infertility. To investigate this, a model of mitochondrial dysfunction was developed in mouse oocytes using a method of photosensitization of the mitochondrion-specific dye, rhodamine-123. After in vitro fertilization, dye-loaded and photosensitized oocytes showed developmental arrest in proportion to irradiation time. Morphological and metabolic assessments of zygotes indicated an increase in mitochondrial permeability that subsequently resulted in apoptotic degeneration. Development was partially restored by inhibition of mitochondrial permeability transition pore formation by oocyte pretreatment with cyclosporin A. Oocyte mitochondria are therefore physiological regulators of early embryo development and potential sites of pathological insult that may perturb oocyte and subsequent preimplantation embryo viability. These findings have important implications for the treatment of clinically infertile women using assisted reproductive technologies.

apoptosis, assisted reproductive technology, embryo, in vitro fertilization, oocyte development


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Mitochondrial dysfunction has been well documented in association with reduced male fertility, but there is surprisingly little equivalent evidence regarding the oocyte, the progenitor cell of the developing embryo [1]. Such information would be valuable for clinical in vitro fertilization (IVF) programs where, a) there are currently few routinely used functional markers of oocyte viability other than cellular morphology and b) reproductive outcomes attributed to reduced oocyte viability remain largely unexplained.

Mitochondria represent a ubiquitous population of bacterium-like structures that are specialized to the production of chemical energy by respiration. Proton and charge gradients inside the mitochondria drive the oxygen-dependent production of energy-rich adenosine triphosphate (ATP) from cytoplasmic energy substrates. Mitochondria also act as stores of intracellular calcium and proapoptotic factors. While predominantly under nuclear control, mitochondrial replication is uniquely self-regulated by multiple copies of mitochondrial DNA (mtDNA) as well as intrinsic protein synthetic and transport machinery.

Mature mammalian oocytes are maternally endowed with thousands of nonreplicating mitochondria that act as the founding population of all daughter-cell mitochondria of the developing embryo [2–5]. Importantly, oocyte mitochondria are relatively devoid of mtDNA templates, the result of a programmed selection process called a genetic bottleneck. This evolutionary process has been proposed to be responsible for the exclusion of mutated mtDNA templates from the germline, but may render oocytes more susceptible to mutagenic mtDNA damage due to lack of redundant templates or active repair mechanisms [6, 7]. Other differences, such as reduced organelle size and less complex internal structure, typify oocyte mitochondria as morphologically primitive or immature compared with those of somatic cells [8, 9]. In spite of this immaturity, metabolic evidence suggests that the mitochondria of the oocyte and early embryo are constitutively active and maintenance of this low-level activity is necessary for ongoing development. Oxygen consumption is low in oocytes and cleavage-stage embryos; however, exclusion or oversupply of oxygen results in embryonic arrest in culture [8–11]. Partial downregulation of aerobic metabolism results in a shift to less efficient anaerobic energy production, causing overproduction of lactate and consequently reduced embryo viability [12–14]. Transient chemical inhibition of oxidative phosphorylation can also induce alterations in preimplantation embryo development in a dose- and stage-dependent manner [9, 15]. Below a particular threshold, mitochondrial ATP has been correlated with reduced developmental competence and postimplantation outcomes [16].

It is therefore a sensitivity of oocyte mitochondria to physiological insults that may contribute to the distinctive susceptibility of the oocyte to developmental compromise, particularly in vitro, and that is perhaps exacerbated by maternal inheritance and a delay in active mitochondrial replication until postimplantation development [2, 17]. To explore this hypothesis, a method of mitochondrial photosensitization previously used as a mitochondrion-specific but not metabolic pathway-specific inducer of mitochondrial damage and subsequent cytotoxicity [18, 19] was applied to a standard mouse model of IVF and preimplantation embryo culture. Mitochondrion-specific fluorophores have been successfully used to study the dynamics of mitochondrial patterning in oocytes and preimplantation embryos [13]. Prolonged excitation of loaded mitochondrial fluorophores (e.g., Mitotracker RedTM), using similar loading protocols, can induce intramitochondrial oxidative stress that damages mitochondria and causes them to leak and set off a cascade of apoptotic degeneration [20]. In the present study, metaphase II mouse oocytes were loaded with rhodamine-123 (R123; Molecular Probes Inc., Eugene, OR) and photosensitized for variable time intervals to induce mitochondrial damage. Endpoints of blastocyst development, zygote mitochondrial function, and zygote apoptosis after standard IVF were subsequently performed.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animals

In all experiments, C57BL6 x CBA/Ca F1 hybrid mice were used. Mice were housed in an environmentally controlled room at 22–24°C with a 12L:12D photoperiod with food and water available ad libitum. For oocyte harvest, 4–6 wk (pubertal) female mice were superovulated by intraperitoneal (i.p.) injection of 5 IU eCG (Folligon; Intervet) followed approximately 48 h later with an i.p. injection of 5 IU hCG (Chorulon; Intervet). For in vitro insemination of oocytes, mature sperm were isolated from whole epididymes of sexually mature (10–12 wk) male mice. Both female and male mice were killed by cervical dislocation before gamete removal.

Ethics of Experimentation

The present study was approved by Monash Medical Centre Animal Ethics Committee A, Monash Medical Centre, Clayton, Victoria, Australia, approval numbers A2003/40. Experiments were conducted in accordance with the 1997 NH and MRC Australian Code of Practice for the Care and Use of Animals for Scientific Purposes and the Victorian Prevention of Cruelty to Animals Act and Regulations, 1986.

Oocyte Isolation, In Vitro Fertilization, and Blastocyst Culture

Oocyte collection, sperm preparation, and in vitro fertilization were performed as described previously [21]. Normally fertilized zygotes were then cultured for 96 h to the blastocyst stage in a humidified atmosphere of 5% CO2 at 37°C. Zygotes were cultured in groups of 10 in 20-µl droplets of bicarbonate-buffered potassium simplex optimized medium with amino acids (KSOMaa) [22] supplemented with 1 mg/ml bovine serum albumin (BSA) (Invitrogen) overlayed with mineral oil (Sigma Aldrich). Blastocysts were cultured in three replicate experimental groups of at least 30 zygotes and formation was expressed as a percentage of normally fertilized oocytes.

Photosensitization of Oocytes Using Epifluorescence Microscopy

Gross mitochondrial dysfunction rather than antagonism of a specific mitochondrial biochemical pathway was induced as described in a published protocol [23], with some modifications. Oocytes were stained for 10 min in HEPES KSOMaa supplemented with 50 µg/ml rhodamine 123 (R123; Molecular Probes) prepared from a stock solution of 100 mg/ml R123 in DMSO (Sigma Aldrich). Loaded oocytes were rinsed twice in HEPES KSOMaa, stored in drops of the same medium on an inverted epifluorescence microscope stage and irradiated with focused visible light generated from a 100-W mercury bulb and filtered through a dichroic mirror (480 ± 10 nm) to excite R123. Test oocytes were irradiated for periods of 0, 20, 40, or 60 sec before insemination. Control oocytes from the same batch of mice were allocated to one of three groups: (a) not loaded with R123 and irradiated for the maximal time period of 60 sec (light), (b) incubated in R123 vehicle DMSO for 15 min before insemination, but not loaded or irradiated (vehicle), or (c) not pretreated, loaded, or irradiated (no treatment).

Laser-Scanning Confocal Microscopic Visualization and Determination of Mitochondrial Membrane Potential ({Delta}{psi})

Assessment of mitochondrial {Delta}{psi} was performed according to the method described by Wilding and colleagues [46]. Zygotes were stained approximately 9 h after fertilization with 20 µg/ml 5,5',6,6'-tetrachloro-1,1,3,3'-tetraethylbenzimdazoylcarbocyanine iodide (JC-1) (Molecular Probes). Individual photosensitized and nonphotosensitized control oocytes were mounted in HEPES KSOMaa and visualized using a krypton/ argon laser scanning confocal microscope (LSCM) (Bio-Rad) configured to detect 520-nm (green) and 580-nm (red) excitation wavelengths. Images of individual zygotes were captured and quantitated for fluorescence intensity using image-analysis software (Scion Image). A test group of at least 20 zygotes formed from loaded oocytes that were irradiated for 60 sec was used, compared with a similar sized control group of zygotes that had fertilized normally from loaded, nonirradiated oocytes. LSCM was also used to capture equatorial sections of zygotes derived from loaded (irradiated or nonirradiated) oocytes for a qualitative visual assessment of oocyte cytoplasmic distribution of R123 (520-nm emission).

Fluorometric Determination of Cytoplasmic NADH-NADPH

Quantitative assessment of autofluorescence at 340-nm excitation (ultraviolet light) was performed according to the microfluorometric method described previously [12]. In this study, values of whole zygote autofluorescence were used as an indicator of cytoplasmic NADH-NADPH content. Test and control groups of zygotes selected were as for those described for determination of {Delta}{psi}, using a separate group of zygotes. Absolute values of autofluorescence intensity were expressed in arbitrary values on a scale of 0–850. All values were corrected for background fluorescence and internal reflectance.

Fluorometric Determination of ATP Content

Cytoplasmic ATP content was performed using a modification of the luciferin/luciferase method previously employed for oocytes and preimplantation embryos [24]. Reference solutions of 0–0.1 mM ATP (Sigma Aldrich) in ultrapure water were prepared for standard curve generation. A commercial luciferin/luciferase reaction mixture (Molecular Probes) was diluted 1:3 in ultrapure water before use. Microdrops of approximately 1 µl of this reaction mixture were positioned in glass-bottomed Petri dishes (Intracel) overlaid with mineral oil. From these drops, smaller drops of 10 nl were aliquoted using a manual micromanipulator and calibrated constriction pipettes [12]. Zygotes were stripped of zona pellucidae by incubation in warmed HEPES KSOMaa containing 0.2% protease (Sigma Aldrich) for approximately 5 min. Groups of 10 zona-free oocytes were then rinsed twice in HEPES KSOMaa, transferred to 250-nl microdrops of ultrapure water and lysed mechanically using a narrow bore hand-drawn glass pipette. From the 250-nl test drops, 1-nl aliquots were added to the 10-nl aliquots of reaction mixture. At the same time, five 1-nl aliquots were taken from each 1-µl ATP standard drop and added to five individual 10-nl reaction drops. Fluorometry was performed as for autofluorescence measurements to determine the decrease in fluorescence of luciferin at 340-nm excitation as ATP was consumed. Test and controls were as for those described for determination of {Delta}{psi} using a separate group of zygotes, although sample numbers represent pooled batches rather than single zygotes.

Assessment of Apoptosis

Propidium iodide staining of chromatin was performed by incubation of zygotes derived from treated oocytes for 15 min in warmed HEPES KSOMaa containing 20 µg/ml of propidium iodide (Sigma Aldrich). Zygotes were then scanned using LSCM set to 490-nm excitation and 580-nm (red) emission set at higher sensitivity, to detect low-level fluorescence. Propidium iodide penetration was also used as an indicator of plasma-membrane damage that occurs in late-apoptotic/early necrotic cells [25]. Detection of caspase activation was performed using a previously described fixation and staining protocol [26]. Exceptions to the protocol include the use of a rabbit monoclonal antibody raised against cleaved mouse caspase-3 (Cell Signalling) as the primary antibody and Texas Red conjugated goat polyclonal IgG (Molecular Probes) as the secondary antibody.

Treatment of Photosensitized Oocytes with Cyclosporin A

Immediately before photosensitization at the maximal exposure time of 60 sec, a separate group of oocytes were preincubated for up to 10 min in a solution of HEPES KSOMaa containing 40 µg/ml cyclosporin A (CsA) (Sigma Aldrich). These oocytes were then inseminated and cultured to the blastocyst stage alongside control zygotes photosensitized in the absence of exposure to CsA.

Statistical Analysis

The mean proportion of zygotes developing to blastocyst by 96 h in vitro was calculated for test and control treatments and exponentially transformed for comparison using ANOVA. Mean values for metabolic measurements and the proportion of zygotes developing to blastocyst following exposure to CsA were compared with control values using chi-squared analysis. Differences between means were considered statistically significant at a P-value of less than 0.05.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Blastocyst Formation Following Oocyte Photosensitization

Of those inseminated oocytes that were loaded and irradiated for 20, 40, or 60 sec, 64%, 52%, and 28% developed to blastocysts (Fig. 1). The latter two proportions are significantly lower (P < 0.01) than those of blastocysts that developed from control groups of oocytes that had been loaded but not irradiated (73%), vehicle treated (77%), irradiated but not loaded (79%), or untreated (84%) (Fig. 1). The relationship of blastocyst development to irradiation time represented a negative trend that correlates exactly (R2 = 1) with the second-order polynomial function y = –0.0075x2– 0.35x + 73, where y represents blastocyst development and x represents photosensitization duration. Importantly, there was no significant decrease in blastocyst development induced by probe loading without irradiation. The majority of embryonic arrest was observed between the zygote and two-cell stages of development.



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  		FIG. 1. Blastocyst development after oocyte photosensitization. Graph of mean blastocyst formation rates of zygotes derived from oocytes photosensitized (lined bars) for 0 sec (n = 100), 20 sec (n = 127), 40 sec (n = 100), and 60 sec (n = 156). Control groups (white bars) represent oocytes that were (a) not loaded but maximally irradiated (light, 60 sec, n = 100), (b) incubated briefly in the R123 diluents DMSO (vehicle, n = 100), or (c) not treated (no treatment, n = 100). Bars sharing like superscripts (1, 2) are not statistically different

Mitochondrial Morphology and Metabolic Function in Zygotes

LSCM analysis of zygotes derived from oocytes loaded with R123 and photosensitized for 60 sec (maximal photosensitization) revealed a diffuse, uniform ooplasmic staining pattern with a defined boundary at the oolemma (Fig. 2A). Zygotes developing from loaded nonirradiated control oocytes, however, exhibited a uniform punctate staining pattern, which is typical of functional, intact mitochondria (Fig. 2B). The diffuse pattern was interpreted as a release of mitochondrial R123 back into the ooplasm and hence evidence of a mitochondrial permeability transition (mPT). Occasionally, plasma membrane permeabilization and large-scale leakage of R123 from the ooplasm itself was observed (unpublished data). None of these changes were observed in untreated or loaded nonirradiated controls, even though R123 is to some degree intrinsically cytotoxic without photosensitization [27].



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  		FIG. 2. LSCM evidence of mitochondrial damage after oocyte photosensitization. Images of mouse zygotes represent equatorial sections taken approximately 9 h after insemination. A) Zygote derived from an oocyte loaded with R123 and maximally photosensitized showing a diffuse staining pattern. B) Zygote derived from R123-loaded nonirradiated oocyte showing a uniform punctate mitochondrial staining pattern. C) Propidium iodide-stained zygote derived from a maximally irradiated oocyte. D–F) Caspase-3 antibody staining of zygotes derived from untreated, loaded nonirradiated, and maximally photosensitized oocytes, respectively. G, H) Arrested zygote and two-cell embryo derived from maximal photosensitization. Note the more pronounced and irregular staining pattern compared with Figure 2A. I) Zygote derived from maximally irradiated oocytes preloaded with CsA showing partial punctate morphology and partial diffuse morphology. J) JC-1 stained zygotes derived from maximally photosensitized oocytes pretreated with CsA (upper image: zygote with active membrane potential; lower image: zygote with reduced or inhibited membrane potential. V, Vacuolation; PB, polar body; PN, pronucleus; scale bar = 10 µm

Quantitative LSCM analysis of zygotes derived from maximally irradiated oocytes revealed a significantly lower {Delta}{psi} (Table 1) compared with loaded nonirradiated control zygotes (8.2 ± 0.33 vs. 4.27 ± 0.30 arbitrary units, P < 0.0001). Microfluorometric analysis revealed that zygotic pools of NADH-NADPH after loading and maximal irradiation were significantly lower (P < 0.05) than after loading without irradiation (158 ± 22 vs. 125 ± 10 arbitrary units), as was zygote ATP content (3.06 ± 0.79 vs. 1.0 ± 0.1 pmol/zygote). These changes to zygote metabolism were interpreted as further evidence of biochemical dysfunction of mitochondria resulting from mPT and metabolic uncoupling.


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  		TABLE 1. Effect of oocyte photosensitization on endpoint measures of zygote mitochondrial physiology.a

Evidence of Mitochondrial Induction of Apoptosis

LSCM analysis revealed low-level penetration of propidium iodide into (unfixed) zygotes derived from maximally photosensitized oocytes (Fig. 2C), which is indicative of a late apoptotic or a necrotic phenotype. An apoptotic phenotype was confirmed by the presentation of male and female pronuclei, in a representative section of a propidium iodide-stained zygote, as dense bodies of compacted chromatin (Fig. 2C) [25], which is different from the expected dispersed disc-like morphology of pronuclear chromatin in nonapoptotic mouse zygotes [28]. Apoptosis was also confirmed by a more intense staining of activated ooplasmic caspase-3 in fixed and antibody-labeled zygotes derived from maximally photosensitized oocytes, in comparison with loaded nonirradiated and untreated control zygotes (Fig. 2, D–F). Furthermore, arrested zygotes and early embryos derived from maximally photosensitized oocytes showed a more pronounced and irregular pattern of mitochondrial R123 staining, which was interpreted as evidence of either aggregation or severe swelling of mitochondria (Fig. 2, G and H).

Pretreatment of maximally photosensitized oocytes with CsA resulted in a significant improvement in blastocyst development after IVF and culture, compared with control oocytes without pretreatment (51% vs. 28%, P < 0.0001) (Fig. 3). This developmental rescue of zygotes derived from oocytes pretreated with CsA was confirmed by partial punctate R123 staining, indicative of restoration or maintenance of intact mitochondria (Fig. 2I) as well as intense red JC-1 staining indicative of maintenance or restoration of {Delta}{psi} (Fig. 2J).



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  		FIG. 3. Improvement in blastocyst development following oocyte photosensitization by stabilization of membrane permeability. Blastocyst development after fertilization of oocytes loaded with rhodamine 123 (R123) and maximally photosensitized for 60 sec (60). The white bar represents oocytes that had been preincubated for 10 min in handling medium containing CsA (+CsA) before photosensitization and the lined bar represents oocytes that were not exposed to CsA (–CsA) (*, P < 0.0001)


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Mitochondrial dysfunction in oocytes was found to be directly responsible for the early arrest of preimplantation embryos in vitro. Prolonged photosensitization of oocytes loaded with R123 triggered morphological and metabolic damage indicative of zygote mitochondrial dysfunction, followed by apoptotic degeneration.

Photosensitizers have been used widely as agents of cytoplasmic- and organelle-specific cytotoxicity with the degree related to the duration of photoirradiation [29]. Acute cytotoxicity following photosensitization of cancer cell lines loaded with R123 has been shown to result in gross and irreversible structural damage to mitochondria [19]. A maximal oocyte photosensitization time of 60 sec in the present study resulted in severely inhibited blastocyst development after fertilization, with a low level blastocyst survival that was possibly due to oocyte-specific variations in numbers of mitochondria that resisted damage. The same maximal treatment of oocytes also resulted in a decreased mitochondrial membrane potential in zygotes, compared with those derived from loaded nonirradiated oocytes. Taken together with the morphological evidence, this loss of membrane potential is suggestive of a mPT [30].

One reported metabolic consequence of sustained mPT is a phenomenon referred to as mitochondrial uncoupling [9]. Normally, mitochondria use a combined electron and proton gradient across a patent inner mitochondrial membrane to drive ATP synthesis and export. However, mitochondria become uncoupled when the proton gradient is abolished or the patency of the inner membrane is lost, but oxygen-dependent electron transfer continues to deplete matrix and cytoplasmic pools of NADH, NADPH, and other electron carriers. Indeed, zygotic pools of autofluorescent NADH/NADPH were significantly depleted after maximal photosensitization treatment compared with nonirradiated controls.

Uncoupled mitochondria are also characterized by reduced or abolished ATP synthesis due to decreased inner membrane integrity and proton gradient. Zygotes derived from maximally photosensitized oocytes contained significantly lower cytoplasmic ATP levels compared with loaded nonirradiated controls. ATP values of zygotes derived from maximally photosensitized oocytes were lower than the threshold value of 2.0 pmol in human oocytes, above which a correlation with increased likelihood of normal early embryo development and implantation after fertilization has been made [16]. Therefore, the reduction in ATP levels may represent a direct limiting of energy available for cellular processes. Raised ATP levels, however, have also been shown to be associated with reduced viability of mouse oocytes [31]. It has also been demonstrated that treatment of later stage bovine preimplantation embryos with low doses of the chemical uncoupling agents sodium azide and 2,4-dinitrophenol improved blastocyst formation and quality in vitro [9], perhaps due to counteraction of already raised levels of ATP. Further experiments would be needed to determine the utilization of zygote ATP (as well as ADP) after R123 photosensitization.

The potential for oocyte mitochondria to mediate or even initiate apoptotic degeneration in oocytes and preimplantation embryos is feasible because oocytes and zygotes express a variety of genes that encode pro- and antiapoptotic proteins [32]. Furthermore, extracellular inducers of apoptosis in somatic cells such as growth factors (e.g., TNF-alpha) [33] or chemical agents (e.g., hydrogen peroxide) [34] are able to induce apoptotic phenotypes in oocytes and preimplantation embryos. In somatic cells, intracellular triggers such as reactive oxygen species (e.g., superoxide and hydroxyl radicals) have been reported to accompany the formation of mPT pores that allow the release of proapoptotic factors (e.g., cytochrome-c) and in turn activate effector caspases that signal genetic and cellular degradation [35, 36]. Photosensitization of mitochondrially loaded R123 in cultured hepatocytes has been reported to react directly with NADPH in the presence of oxygen to produce singlet oxygen, a transient but highly reactive oxygen species [37, 38]. Singlet oxygen is capable of nonspecific oxidation of lipids, protein, and chromatin, and its production has been reported to coincide with opening of mPTs before the onset of apoptotic cell death [7, 37–39].

As an initial test for the occurrence of mitochondrion-triggered apoptosis, zygotes derived from maximally photosensitized oocytes were stained with the DNA fluorophore propidium iodide. LSCM analysis revealed low-level zygote penetration of propidium iodide, which was interpreted as plasma membrane damage associated with a late-apoptotic/early-necrotic phenotype. The appearance of compacted rather than dispersed male and female pronuclear chromatin was, however, more indicative of late-apoptotic DNA damage [25]. A second confirmation of an apoptotic transition as a consequence of a mPT was a higher fluorescence intensity of activated cytoplasmic caspase-3 in fixed and antibody-labeled zygotes derived from maximally photosensitized oocytes, compared with zygotes raised from untreated and loaded nonirradiated control oocytes. Positive immunohistochemical staining of active caspases has previously been reported in association with chemically induced apoptosis in mouse zygotes [26]. Caspase-3 is the key effector caspase in somatic cells and has been reported previously to be important in apoptosis of mouse oocytes in vivo [40]. Similar to the previously described swelling and aggregation patterns of apoptotic phenotypes [41], the more pronounced and irregular pattern of mitochondrial granularity of arrested zygotes and early embryos was taken as a third indicator of apoptosis.

Transient exposure of oocytes to CsA, a chemical inhibitor of mPT pore formation, [30] before R123 loading and maximal photosensitization resulted in a significant improvement in blastocyst development after insemination compared with unexposed oocytes. Zygotes derived from oocytes treated with CsA also exhibited a restoration or maintenance of mitochondrial morphology and function. CsA in previous somatic cell studies has been shown to react with cyclophilin, a mitochondrial protein that participates in mPT pore formation in association with apoptosis [42]. The rescue of preimplantation development following photosensitization provides further evidence that the developmental failure observed is due to the mPT, and more specifically the formation of mPT pores, which promote the release of proapoptotic factors [43] and recruit proteins such as Bcl-2, as reported in processes of oocyte apoptosis [44]. Another mPT pore inhibitor, bongkrekic acid, has recently been shown to lead to a reduction in ATP content of mature mouse oocytes without detriment to mitochondrial membrane potential [45]. This modulation of ATP levels was attributed to inhibitory effects of bongkrekic acid on adenine nucleotide translocator, a protein responsible for ATP export from mitochondria as well as a participant in mPT pore formation. This result confirms that oocyte mitochondria, like somatic cell mitochondria, are indeed capable of mPT pore formation in response to external stimuli.

In summary, the present study provides new evidence that oocyte mitochondria make a necessary physiological contribution to the cytoplasmic regulation of preimplantation embryo development. The data provide a functional link between a previously reported age-related reduction in human oocyte mitochondrial membrane potential (Wilding et al. [46]) and an age-related decline in human preimplantation embryo developmental potential [46, 47] in association with female infertility. Mitochondrial pathophysiology of the oocyte has, however, not yet been directly identified as a causal factor in female infertility, so further assessments of mitochondrial function in human oocytes are required to answer this basic question. Relevant and potentially causally linked factors include age-related increases in the incidence of apoptosis and chromosomal defects [48, 49], mtDNA defects [50], and deficits in mitochondrial gene expression [51] in human oocytes from infertile patients.

Mitochondrial photosensitization using R123 in the mouse or other appropriate animal models may aid in the design of ART (assisted reproductive technologies) strategies, at the single oocyte level, that will allow for prediction and restoration of compromised developmental competence in clinically infertile women.


   ACKNOWLEDGMENTS
 
This study was undertaken as part of research toward a Ph.D. The authors wish to thank Dr. Peter Crack for his donation of the caspase-3 antibody.


   FOOTNOTES
 
1 Supported by departmental funding based on private donation. Back

2 Correspondence: George A. Thouas, Monash Institute of Reproduction and Development, Monash Medical Center, 246 Clayton Rd., Clayton, Victoria 3168, Australia. FAX: 61 3 9594 7311/7314; george.thouas{at}med.monash.edu.au Back

Received: 23 June 2004.

First decision: 12 July 2004.

Accepted: 22 July 2004.


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 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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