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RESEARCH |
Department of Equine Sciences, Faculty of Veterinary Medicine, Utrecht University, Yalelaan 12, 3584 CM Utrecht, The Netherlands, 1 Laboratorio di Tecnologie della Riproduzione (LTR-CIZ), Istituto sperimentale Italiano Lazzaro Spallanzani, Via Porcellasco 7/F, Cremona 26100, Italy and 2 Dipartimento Clinico Veterinario, University of Bologna, V Tolara di Sopra 50, Ozzano Emilia, Bologna, Italy
Correspondence should be addressed to T A E Stout; Email: t.a.e.stout{at}vet.uu.nl
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Introduction |
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At recovery from an ovarian follicle, a normal oocyte is surrounded by a corona radiata and a number of layers of cumulus cells. These ‘cumulus oocyte complexes’ (COCs) are generally classified as being either compact (Cp) or expanded (Ex), depending on the degree of cumulus expansion (Hinrichs et al. 1993). Immature oocytes with an Ex cumulus have already embarked on cytoplasmic maturation and therefore require less time to synthesize the proteins needed to resume meiosis (Alm & Hinrichs 1996); they consequently resume meiosis and reach MII in vitro faster than immature oocytes from Cp COCs (Hinrichs et al. 1993). In cattle, the majority of COCs recovered from slaughterhouse ovaries have a Cp cumulus investment (>80%; Lonergan et al. 1994) and, in most laboratories, only Cp COCs are used for in vitro maturation (IVM)/fertilization because they are a more uniform group with significantly better mean developmental competence (Blondin & Sirard 1995). In horses, a far greater proportion of recovered COCs are Ex or partially Ex (approximately 45%: Hinrichs 1997) and, even though Ex COCs are thought to originate predominantly from atretic follicles (Hinrichs & Williams 1997), their rates of IVM (Hinrichs & Williams 1997) and embryo development following ICSI (Tremoleda et al. 2003a, 2003b, Choi et al. 2004) are similar to or better than Cp COCs.
To date, the few studies conducted on the cryopreservability of equine oocytes have concentrated on Cp GV stage COCs, and the success of subsequent IVM has been disappointing (16–30% MII; Hochi et al. 1994, 1996, Hurtt et al. 2000). Nevertheless, two foals were produced following oocyte-transfer of 26 oocytes vitrified after maturation in vivo (Maclellan et al. 2002), thereby demonstrating that in optimal conditions (maturation, fertilization, and embryo development in vivo) equine oocytes can retain developmental competence after vitrification. In a recent study to investigate whether equine oocytes survive freezing better at the GV or at the MII stage, we found that controlled-rate freezing (CF) at the MII stage yielded a higher percentage of MII oocytes with a normal spindle (67%) than CF at the GV stage (1%) or vitrification at either the GV (52%) or the MII stage (37%: Tharasanit et al. 2006). However, the cleavage rate for oocytes fertilized by ICSI after CF at the MII stage was poor (8%), while the ability of vitrified oocytes to support fertilization was not examined. Given the poor developmental competence of CF MII oocytes, it seems logical to investigate now whether vitrifying equine oocytes allows them to maintain developmental competence despite a higher incidence of spindle damage. In addition, since cumulus cells are thought to protect the oocyte against cell damage during cryopreservation (Imoedemhe & Sigue 1992, Pellicer et al. 1988), it is relevant to examine whether Cp or Ex equine COCs are more suited to cryopreservation. Therefore, the present study aimed to examine the effect of maturation stage (GV vs MII) and cumulus morphology at collection (Cp vs Ex) on the ability of equine oocytes to withstand vitrification, in terms of post-thaw maturation rates and MII spindle quality, and via cleavage and embryo development rates following intracytoplasmic sperm injection (ICSI).
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); COCs with fewer cumulus cell layers were discarded. The COC classification was performed using a dissecting stereomicroscope by a single observer to ensure the uniformity of COC types throughout the experiments. They were then maintained at 37 °C in HEPES buffered M199 (Sigma-Aldrich Chemicals BV) supplemented with 0.014% (w/v) BSA (holding medium, HM).
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Open pulled straw (OPS) vitrification of COCs
OPS vitrification was performed essentially as described by Vajta et al.(1998). OPS straws were produced from 0.25 ml polyvinyl chloride straws (IMV technologies; L’Aigle, France) by heating them over a 100 °C hot plate and then stretching until their outer diameter was approximately halved. In preparation for vitrification, groups of ten oocytes were first immersed for 30 s in a 100 µl droplet of HM supplemented with 10% (v/v) ethylene glycol (EG) and 10% (v/v) dimethyl sulfoxide (DMSO; both Sigma-Aldrich Chemicals BV). Next, the oocytes were immersed in a 100 µl droplet of vitrification solution; HM containing 20% EG, 20% DMSO, and 0.5 M sucrose. After 15 s incubation, the COCs were transferred to a 2 µl droplet of fresh vitrification solution and loaded into a modified straw by capillary action. In total, oocytes were exposed to vitrification solution for 20–25 s prior to immersion in liquid nitrogen.
After 3–6 weeks of storage, oocytes were warmed by immersion in a 37 °C ‘warming solution’ consisting of 0.3 M sucrose in HM. The oocytes were then expelled into the warming solution and the cryoprotectant (CPA) was removed by equilibrating the oocytes in this solution for 5 min. Thereafter, the oocytes were washed once and maintained in HM until further examination.
To examine whether CPA was harmful to oocytes, control immature and in vitro matured COCs (both Cp and Ex) were exposed to the vitrification and warming solutions without intervening vitrification.
Experiment I: Effect of cumulus morphology at recovery and maturation stage at vitrification on maturation rate and meiotic spindle quality
COCs classified at recovery as Cp or Ex were vitrified by OPS either immediately (GV stage; n = 63 Cp, 59 Ex) or after 24 h IVM (MII: n = 79 Cp, 83 Ex). Subsequently, COCs vitrified at the GV stage were warmed and incubated in vitro for 30 h, while those vitrified after 24 h IVM were warmed and incubated for a further 6 h, prior to fixation in 4% paraformaldehyde. A further 80 Cp and 83 Ex COCs were used to investigate whether exposure to CPA without vitrification also affected the ability of GV oocytes to resume meiosis, or the quality of the cytoskeleton at the MII stage. Of these, 40 Cp and 42 Ex COCs were exposed to CPA prior to IVM, and the remaining 40 Cp and 41 Ex COCs were first matured in vitro and then treated with CPA. A final 80 Cp and 80 Ex COCs were used as untreated controls. Stage of maturation and cytoskeleton quality of all oocytes were examined by confocal laser scanning microscopy (CLSM) after staining with fluorescent labels for actin microfilaments, microtubules, and chromatin.
Staining the meiotic spindle and chromatin
Prior to staining, cumulus cells were removed from matured oocytes by vortexing the COCs in calcium and magnesium free Earle’s balanced salt solution (EBSS: Gibco BRL, Paisley, UK) containing 0.1% (w/v) hyaluronidase and 0.25% (v/v) trypsin EDTA (both Sigma-Aldrich Chemicals BV) at 37 °C. The denuded oocytes were then incubated for 30 min at 37 °C in a glycerol-based microtubule stabilizing solution containing 25% (v/v) glycerol, 50 mM KCL, 0.5 mM MgCl2, 0.1 mM EDTA, 0.1 mM EGTA, 1 mM 2-mercaptoethanol, 50 mM imidazol, 4% Triton-X-100, and 25 µM phenylmethyl-sulfonyl fluoride (all Sigma-Aldrich Chemicals BV) at pH 6.7 (Simerly & Schatten 1993). After a subsequent wash with PBS supplemented with 0.1% BSA, the oocytes were fixed and stored for 1–2 weeks in 4% (w/v) paraformaldehyde in PBS. To stain the microtubules, oocytes were first incubated for 1 h at room temperature with a 1:100 solution of a monoclonal anti-
-tubulin antibody (clone B1-5-1-2: Sigma-Aldrich Chemicals BV) in PBS–BSA. After a further wash with PBS–BSA, the oocytes were incubated for 1 h with a 1:100 solution of a goat anti-mouse secondary antibody conjugated to tetramethylrhodamine isothiocyanate (TRITC) in PBS supplemented with 2% (v/v) goat serum (both Sigma-Aldrich Chemicals BV). Next, the oocytes were incubated in a solution of 0.165 µM Alexa Fluor 488 phalloidin (Molecular Probes Europe BV, Leiden, The Netherlands) in PBS–BSA for 30 min to label the actin microfilaments and, finally, they were incubated for 15 min in 20 µM TO-PRO3 (Molecular Probes) to stain the chromatin.
Confocal laser scanning microscopy (CLSM)
To assess oocyte quality, fluorescently labeled oocytes were mounted on glass microscope slides in a 2 µl droplet of antifade medium (Vectashield, Vector Labs, Burlingame, CA, USA) to retard photobeaching, and sealed under a coverslip using nail polish. Examination was performed using a confocal laser-scanning microscope (Leica TSC MP, Heidelberg, Germany) mounted on an inverted microscope (Leica DM IRBE). Three laser sources (Argon-ion 514 nm, Krypton 563 nm and HeNe 633 nm) were used to simultaneously excite the fluorescent signals from the Alexa Fluor 488 phalloidin (microfilaments), TRITC (microtubules), and TO-PRO3 (DNA) via 488/568/650 nm excitation/barrier filter combinations. Oocytes were examined using the sequential scanning mode so that the spindle could be examined in three dimensions (including the Z plane); digital micrographs produced for the three separate colors were then merged into single panels using Leica confocal software (version 2.61: Leica Microsystems, Heidelberg, Germany). The resulting multi-color micrographs were subsequently examined using Adobe Photoshop 7 (Adobe Systems Inc., San Jose, CA, USA), and oocytes were classified by the normality of their cytoskeleton, as described earlier by Saunders & Parks (1999). In short, an oocyte’s actin cytoskeleton was classified as normal if the microfilaments were distributed evenly throughout the tranzonal channels and ooplasm, except for a slight intensification just inside the oolema. The microtubulin meiotic spindle was classified as normal if it was symmetrically barrel-shaped with two anastral poles and two equal sets of chromosomes aligned at its center (Fig. 1C
), as described by Tremoleda et al.(2001). Any changes in the distribution of the actin or in the appearance of the meiotic spindle were classified as minor or severe abnormalities; these included clumping of the actin microfilaments and disorganization or dispersal of microtubules and/or chromatin from the spindle (Fig. 1D). In some cases, oocytes lacked any visible microtubular structures around their sets of chromatin and were therefore recorded to have lost their meiotic spindle. Occasionally, the meiotic spindle was aligned in such a way that it was not possible to properly evaluate its normality; in such cases spindle morphology was classified as unidentifiable (Fig. 1E).
Experiment II: The effect of cumulus morphology at recovery and maturation stage at vitrification on developmental competence of oocytes fertilized by ICSI
After recovery, 685 COCs classified as either Cp or Ex were vitrified by OPS immediately (GV stage: n = 229 Cp, 153 Ex) or after 28–30 h of IVM (MII stage: n = 150 Cp, 153 Ex). To control the possible toxic effects of CPA, additional COCs were exposed to the vitrification media either before (n = 48 Cp, 37 Ex) or after (n = 48 Cp, 47 Ex) IVM, without vitrification. A final 158 Cp and 130 Ex COCs were used as non-treated/non-frozen controls. After warming and CPA removal, oocytes vitrified at the GV stage were matured in vitro for 28 h. For all treatment groups, oocytes that reached MII were fertilized by ICSI and then cultured in vitro, as described later. The cleavage and blastocyst formation rates were recorded 2 and 9 days later respectively. On day 9, the embryos were stained for 10 min at 37 °C with 2 µM Ethidium homodimer-1 (Ethd-1; Molecular Probes) to identify dead cells before being fixed in 4% paraformaldehyde in preparation for further staining.
Intracytoplasmic sperm injection (ICSI)
Matured oocytes designated for fertilization were partially denuded of cumulus cells by incubation in HEPES-buffered synthetic oviductal fluid (H-SOF: Lazzari et al. 2002) containing 25 µg/ml hyaluronidase (Sigma-Aldrich Chemicals BV) followed by aspiration through a pipette tip. Remaining cumulus cells were removed by incubating the oocytes for 1.5 min in a 0.25% (w/v) solution of trypsin (Sigma-Aldrich Chemicals BV) in H-SOF before transfer to H-SOF supplemented with 10% heat-inactivated fetal calf serum (Sigma-Aldrich Chemicals BV) and repeated aspiration through a fine glass pipette. Oocytes with a normal MII appearance, including an extruded first polar body (PB), were considered suitable for ICSI with frozen–thawed sperm from a single fertile stallion.
After thawing, the sperm was rinsed free of cryoprotectant by centrifugation at 750 g through a discontinuous 45:90% Percoll density gradient (Sigma-Aldrich Chemicals BV) at room temperature. Next, the sperm pellet was resuspended in 2 ml Ca2+ free Tyrode’s albumen lactate pyruvate (TALP) (Parrish et al. 1988) and centrifuged at 400 g for 10 min before further resuspension at a concentration of 4 million sperm/ml in SOF supplemented with 6 mg/ml fatty-acid-free BSA (FAF-BSA; Sigma-Aldrich Chemicals BV), modified Eagle’s medium (MEM) amino acids, 1 µg/ml heparin, 20 µM penicillamine, 1 µM epinephrine, and 10 µM hypothaurine (SOF-IVF: Lazzari et al. 2002: all ingredients from Sigma-Aldrich Chemicals BV). Finally, the sperm suspension was diluted 1:1 (v/v) with a 12% solution of polyvinylpyrolidone (PVP; Sigma-Aldrich Chemicals BV) in H-SOF-medium.
Sperm injection was performed as described by Kimura & Yanagimachi (1995). Pipettes produced using a glass micropipette puller (Model P-87, Sutter Instruments, Novato, CA, USA) and with inner diameters of 30 and 5 µm were used for holding oocytes and for sperm injection respectively. ICSI was performed at 37 °C using a micromanipulator (Narishige Co. Ltd, Tokyo, Japan) equipped with a Piezo micropipette-driving unit (Prima Tech, Ibaraki, Japan) and mounted on an inverted microscope (Nikon TE 300: Nikon, Kawasaki, Japan). A motile sperm was immobilized by applying two or three piezo-pulses to its tail-midpiece region, and it was then aspirated into the tip of the injection needle. The oocyte for injection was immobilized using the holding pipette and orientated with its PB at 0600 or 1200 h. The ICSI needle was then advanced through the zona pellucida and oolema at 1500 h using the piezo-drilling motion, and the sperm was released into the ooplasm.
In vitro zygote/embryo culture
Following ICSI, groups of ten sperm-injected oocytes were cultured in 20 µl droplets of SOF-IVC supplemented with MEM amino acids and 16 mg/ml FAF-BSA (SOF-BSA-aa; Galli et al. 2001) under mineral oil at 38.7 °C in an atmosphere of 5% CO2, 5% O2, and 90% N2. On day 2 after ICSI, the presumptive zygotes were examined for cleavage; cleaved embryos were cultured for a further 7 days, during which the culture medium (SOF-BSA-aa) was refreshed on day 4 with SOF by adding 20 µl of new medium and removing 20 µl of the mix. On day 6, the media was again refreshed, this time with DMEM/F-12 (Sigma-Aldrich Chemicals BV) supplemented with 5% (v/v) heat-inactivated fetal calf serum and 5% (v/v) serum replacement (SR: Irvine Scientific, Santa Ana, CA, USA). Finally, on day 9 dead cells in developing embryos were stained with Ethd-1 before fixing the embryo in 4% paraformaldehyde. The labeled embryos were then stored in fixative in the dark at 4 °C for approximately 2–3 weeks until embryo quality was examined.
Assessing the quality of in vitro produced blastocysts
Blastocyst quality was quantified in terms of the percentages of dead cells and of cells with fragmented nuclei or fragmented DNA, as described by Pomar et al.(2005). To identify fragmented DNA, blastocysts previously stained with Ethd-1 were permeabilized in 0.1% Triton X-100 in PBS for 15 min at 4 °C, then washed in PBS–BSA and stained for 1 h at 37 °C with fluorescent-conjugated dUTP and TdT (TUNEL reagent; Boehringer Mannheim, Roche Diagnostics GmbH, Mannheim, Germany). To visualize intact and fragmented cell nuclei and enable calculation of the percentages of dead or TUNEL-positive cells, the blastocysts’ nuclear DNA was stained for 15 min with 4'6-diamidino-2-phenylindole dihydrochloride (DAPI; Sigma-Aldrich Chemicals BV) in PBS–BSA. Once labeled, embryos were mounted in a 2 µl droplet of antifade medium on a glass microscope slide and examined using a conventional fluorescent microscope (BH2-RFCA; Olympus, Tokyo, Japan) using 488, 568, and 350 nm filters for TUNEL, Ethd-1, and DAPI respectively. Counting of blastocyst nuclei was facilitated using an eyepiece counting grid, and representative images were recorded using a digital camera (Coolpix 990, Nikon, Melville, NY, USA).
Statistical analysis
Statistical analysis was performed using SPSS 12.0.1 for Windows (SPSS Inc., Chicago, IL, USA) and logistic regression analysis (MaCullagh & Nelder 1989). Differences between experimental groups in maturation rates, meiotic spindle quality (Experiment I), cleavage, and blastocyst formation rates (Experiment II) were compared using the model: ln 
/(1–) = + treatment, where = frequency of positive outcome, and = the intercept. The total numbers of nuclei per blastocyst (Experiment II) were compared using one-way ANOVA. In all cases, differences were considered significant when P<0.05.
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). For Cp COCs, this maturation percentage did not differ significantly to that for control non-frozen oocytes (58%). By contrast, the maturation rate for vitrified Ex COCs was lower than for non-frozen oocytes (68%), and the reduced success appeared to be due predominantly to a greater incidence of oocytes that resumed meiosis but subsequently arrested in MI (22 vs 4% for vitrified vs control Ex COCs). The salient factor with regard to the reduced ability of Ex COCs to successfully complete meiosis appeared to be vitrification per se, since exposure to CPA without vitrification resulted in maturation rates comparable with non-frozen controls (62%; Table 1).
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). By contrast, when Cp and Ex COCs were first matured in vitro and then vitrified, there was no discernible effect of initial cumulus morphology on meiotic spindle quality, which was poor in both groups (32 and 28% normal spindles for Cp and Ex COCs respectively). In all cases, the average meiotic spindle quality of vitrified–warmed oocytes was worse than in non-frozen controls (95% normal; P<0.05; Table 2), while exposure to CPA had no apparent effect on meiotic spindle quality for either type of COC at either stage of maturation (93–100% normal; Table 2). Although a number of vitrified–warmed oocytes also showed changes from the normal pattern of actin microfilament staining (Tremoleda et al. 2001), such as clumping or very weak staining, these changes were difficult to quantify. Overall, vitrification of Cp COCs followed by IVM yielded a higher percentage of ‘normal’ MII oocytes (product of percentage achieving MII and percentage of the MII oocytes with a normal spindle: 26%) than similar handling of Ex COCs (15%), or IVM followed by vitrification of either type of COC (both 19%).
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). Of the MII oocytes deemed suitable for ICSI, the percentages that had cleaved by day 2 after sperm-injection also did not differ between Cp and Ex oocytes vitrified at the GV stage (34 vs 27% respectively; P>0.05). By contrast, oocytes vitrified after IVM had a lower mean cleavage rate, and Cp COCs fared better than Ex (16 vs 4% cleavage). In all cases except Cp GV, cleavage rates for vitrified COCs were lower than for control oocytes (48 and 66% for control Cp and Ex). Again, the freezing process per se appeared to be the most damaging part of vitrification since cleavage rates for oocytes exposed to CPA either before or after maturation (60–75%; Table 3) were not lower than controls.
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). With regard to blastocyst quality, the mean number of nuclei per blastocyst was 240 ± 18 and neither total cell number nor the percentages of cells that were dead, DNA fragmented or had a fragmented nucleus differed between groups (Table 4; Fig. 1F).
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Although initial cumulus morphology did not affect the proportion of vitrified GV oocytes that reached MII during IVM, it did markedly affect the quality of the MII spindle; fewer oocytes derived from Cp than Ex COCs displayed significant meiotic spindle disruption and chromatin dispersal (37 vs 67% damaged spindles for Cp vs Ex). That oocytes from Ex COCs suffered more spindle disruption than those from Cp COCs may simply be because a tightly compacted cumulus more effectively buffers the oocyte against osmotic damage during vitrification and warming than an Ex cumulus with fewer, or less effective, protective cell layers; Pellicer et al.(1988) earlier reported that oocytes with fewer surrounding cumulus layers were less likely to survive cryopreservation. Moreover, abnormalities of spindle morphology in oocytes matured in vitro after freezing are common (e.g. in man: Van Blerkom & Davis 1994, Park et al. 1997; mouse: Isachenko & Nayudu 1999), and probably relate to freezing-induced impairment of factors critical to the assembly of the meiotic spindle. In this respect, it is also likely that oocytes with an Ex cumulus are more susceptible to damage during vitrification because they are at a more advanced stage of cytoplasmic maturation than those with a Cp cumulus (Alm & Hinrichs 1996). Oocytes with an Ex cumulus at recovery have probably synthesized more of the proteins necessary for completing meiosis and assembling the meiotic spindle (e.g. mitogen activated protein (MAP) kinase: Verlhac et al. 1994), and the mechanisms for spindle assembly and repair may therefore be more susceptible to cryodamage than in oocytes with a Cp cumulus that have yet to embark on synthesis of these crucial proteins.
The alternative to cryopreserving oocytes at the GV stage and maturing them after warming is to mature them first and vitrify them once they have reached MII. Indeed, in an earlier experiment, we achieved the highest proportion of ‘normal’ MII oocytes after CF at the MII stage (35% of oocytes subjected to initial IVM; Tharasanit et al. 2006). Similarly, bovine oocytes have been reported to tolerate chilling and cryopreservation better at the MII than the GV stage (Lim et al. 1992, Parks & Ruffing 1992). Although the reasons for the differences in susceptibility to freezing damage are not known, they probably include factors such as the oolema of MII oocytes being more permeable to water (Ruffing et al. 1993) and cryoprotectants (e.g. DMSO, EG: Agca et al. 1998) and better able to maintain plasma membrane lipid fluidity at low temperatures (Arav et al. 1996). However, in the current study, vitrifying equine oocytes after IVM led to considerable spindle damage (only 28–32% normal spindles), despite a 6 h post-warming incubation to allow spindle repair; in human oocytes, the meiotic spindle disappears completely during freezing but repolymerizes during post-thaw incubation (Rienzi et al. 2004). As a result of the poor spindle quality of oocytes vitrified at the MII stage, the overall proportion of Cp COCs that yielded an MII oocyte with a normal spindle after vitrification tended to be better if they were vitrified at the GV than at the MII stage (26 vs 19%); for Ex COCs, the final percentage of normal MIIs tended to be lower for oocytes vitrified at both the GV and MII stages (15 and 19% respectively). These values are similar to those reported earlier for equine oocytes vitrified at the GV (15%) or MII (20%) stages but, despite an improvement in the percentage of vitrified GV oocytes reaching MII, are still lower than the 35% normal MIIs recorded after CF at the MII stage (Tharasanit et al. 2006). In the latter and the present studies, and in studies on human oocytes (Park et al. 1997, Boiso et al. 2002, Rienzi et al. 2004), the bulk of the damage to meiotic resumption and spindle assembly or morphology during vitrification of horse oocytes was caused by freezing per se, since exposure to CPA without vitrification did not significantly affect either process.
While extensive irreversible damage to the meiotic spindle would severely compromise an oocyte’s ability to be fertilized and give rise to a normal embryo, MII spindle normality alone is not a good indicator of developmental competence because it gives no indication of the state of numerous cytoplasmic factors critical to cell activation during fertilization, and subsequent cell division, fertilization. In this respect, the enzyme cascade that inactivates maturation-promoting factor (MPF) is critical for releasing the oocyte from MII arrest, while inactivation of MAP kinase appears to be involved in pronucleus formation (Sun & Nagai 2003, Fan & Sun 2004). In addition, functional mitochondria are essential for the continuous, active microtubule disassembly and reassembly that is critical to sperm aster formation and syngamy, and mitochondrial damage is a common feature of vitrified–warmed horse oocytes (Hochi et al. 1996). Disruption of mitochondria, enzymes, and structural proteins during vitrification almost certainly explains why the blastocyst formation rate for vitrified–warmed oocytes in the present study was so low (0.4% of injected oocytes compared with 10–17% for controls). Furthermore, since exposure to CPA without freezing had no significant effect on cleavage, blastocyst formation rate (8–12%), or blastocyst quality, freezing per se appears to be the primary cause of reduced developmental competence.
Although it was not possible to compare blastocyst formation rates and quality between groups of vitrified oocytes, there was a marked effect of maturation stage at vitrification on the cleavage rate; oocytes vitrified prior to IVM yielded higher cleavage rates at day 2 after ICSI (34 and 28% of injected oocytes for Cp and Ex respectively) than those vitrified after IVM (16 and 4%); these values were also much higher than for oocytes subjected to CF at the MII stage (8%; Tharasanit et al. 2006). By contrast, while there was no difference in cleavage rates between bovine oocytes vitrified at the GV or MII stage, blastocyst rates were better if maturation had already been initiated (Hochi et al. 1998) or completed (Men et al. 2002, Diez et al. 2005). Therefore, cleavage rate is not a sufficiently reliable indicator of developmental potential. Indeed, while Ex COCs vitrified at the GV stage yielded similar cleavage rates to Cp COCs in the present study, it is likely that their higher incidence of spindle abnormalities would lead to chromosomally abnormal cells and therefore compromise subsequent embryo development; many more blastocysts are needed to test this hypothesis. Although we produced only one blastocyst in the present study (from a Cp COC vitrified at the GV stage), (Maclellan et al. 2002) described the production of three pregnancies from 26 oocytes (12%) matured in vivo, recovered by transvaginal follicle aspiration, vitrified, and subsequently transferred to the oviduct of recipient mares. The superior developmental competence in Maclellan et al.(2002) study was undoubtedly a factor of the superiority of the mare’s follicle and oviduct over in vitro culture systems for oocyte maturation and embryo development. In this respect, oocytes matured in vivo give rise to much better pregnancy rates after transfer to the oviduct of a recipient mare than those matured in vitro (e.g. 82 vs 9%; Scott et al. 2001), while the blastocyst formation rates for equine oocytes matured in vitro and fertilized by ICSI are much better if the presumptive zygotes are transferred to the oviduct of a mare than if they are cultured in vitro (36 vs 0–20%: Choi et al. 2004). Transfer of vitrified–warmed oocytes to the oviduct of recipient mares, after ICSI or insemination of the recipient, would probably therefore have yielded better embryo development rates, and may have offered more information about the effects of cumulus morphology and maturation stage at vitrification on oocyte developmental competence. Nevertheless, from a practical and animal welfare perspective, improving in vitro culture systems is preferable to surgical transfer of oocytes or zygotes to the oviduct.
In conclusion, the present study demonstrated that equine oocytes vitrified at the GV stage can reach MII at respectable rates during post-warming IVM (41–54%). And, while the cumulus morphology at recovery (Cp vs Ex) did not affect the cleavage rates after ICSI (approximately 30% of injected oocytes), it did significantly affect the quality of the meiotic spindle in MII oocytes (63 vs 33% normal spindles for Cp vs Ex COCs). Vitrifying equine oocytes after IVM led to relatively poor rates of both spindle normality (28–32%) and cleavage (4–16%). While CF of MII oocytes may offer an even higher likelihood of a normal looking MII oocyte (Tharasanit et al. 2006), overall it appears that vitrifying equine oocytes with a Cp cumulus at the GV stage probably offers the best prospect for a fertilizable, developmentally competent oocyte. However, since only one blastocyst was produced from 257 sperm-injected vitrified–warmed oocytes, it can presently only be concluded that vitrifying equine oocytes at either stage of maturation leads to a dramatic drop in developmental competence. Further studies are needed to address why vitrified–warmed oocytes that cleaved failed to progress to the blastocyst stage.
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Footnotes |
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References |
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