Sperm and Oocyte Treatments to Improve the Formation of Male and Female Pronuclei and Subsequent Development Following Intracytoplasmic Sperm Injection into Bovine Oocytes¹

^a Departments of Biomedical Sciences ^b and of Population Medicine, Ontario Veterinary College, University of Guelph, Ontario, Canada N1G 2W1

	ABSTRACT

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This study assessed pronuclear formation, the chromosomal constitution, and the developmental capacity of bovine zygotes formed by intracytoplasmic injection of oocytes with sperm, treated or not with dithiothreitol (DTT). Oocytes were matured in vitro for 22–24 h and then centrifuged so that sperm, prepared by swim-up in the presence or absence of 5 mM DTT, could be injected into the cleared area of the ooplasm. Injected oocytes were activated by treatment with 5 µM ionomycin (5 min) and, after a 3-h interval, with 1.9 mM 6-dimethylaminopurine (DMAP) for 3 h. They were then cocultured with bovine oviductal epithelial cells in M199. Sperm treatment resulted in a significantly higher proportion of male pronucleus formation 16 h after injection (40% vs. 11%; p < 0.0001) and a significantly higher rate of blastocyst development (24% vs. 10%; p < 0.005). Sixty-one percent of blastocysts produced with treated sperm were diploid. Of 12 blastocysts produced with treated sperm and sexed by a polymerase chain reaction, 4 were male and 7 female, and in one a definite diagnosis could not be made. Embryo transfer (2 embryos per heifer) resulted in pregnancies in 6 of 16 recipients at Day 49, but none was carried to term. These results show that the efficiency of bovine intracytoplasmic sperm injection can be improved by sperm pretreatment with DTT and by oocyte activation with ionomycin plus DMAP, although the developmental capacity of the resulting embryos remains limited.

	INTRODUCTION

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Since the first report of intracytoplasmic sperm injection (ICSI) in mammals (in the hamster [1]), the transfer of embryos produced by the procedure has given rise to live young in mice [2, 3], rabbits [4], cattle [5], and humans [6–10]. In humans, ICSI has become a very widely applied means of overcoming infertility. In addition to its clinical usefulness, ICSI can be a valuable research tool for studying fundamental aspects of how the two gametes interact during fertilization. For example, in several species the injection procedure itself is apparently sufficient to activate the oocyte, as the sperm nucleus can undergo decondensation and formation of a pronucleus (PN) when injected into the oocyte (humans [6, 9, 11, 12], rabbits [13, 14], hamsters [1, 14], mice [2, 3]). In contrast, ICSI with the bovine spermatozoon rarely leads to its decondensation and male PN formation during subsequent culture in vitro [11, 14–16]. Permeabilization of the sperm membrane by physical means may have a role to play in facilitating decondensation and PN formation after ICSI; artificial removal of the acrosome and tail by sonication [13, 17], immobilizing sperm and damaging the sperm membrane by freezing and thawing before injection [5, 15, 17], and crushing the sperm with the micropipette used for injection [18] have all been reported to improve results.

During the final stages of mammalian sperm maturation, their nuclear structure becomes progressively condensed and stabilized by the formation of disulfide bonds [19–21]. It has also been proposed that decondensation of the sperm nucleus and formation of the male PN are affected by the structural stability of the sperm nucleus [14, 22–25]. This stability depends upon the association of sperm DNA with protamines, the sperm-specific basic proteins that replace somatic histones during spermiogenesis and also render the DNA genetically inactive. Species variation in sperm nuclear stability is therefore considered to account for the ease or difficulty with which sperm decondensation and PN formation occur following ICSI. For example, ICSI into the hamster oocyte readily leads to decondensation of the nucleus of human, mouse, chinchilla, and hamster sperm, but much less easily in the case of rat and bovine sperm nuclei [14]. In fact, bovine sperm nuclei rarely decondense in the hamster oocyte unless they have been treated with dithiothreitol (DTT; an agent that specifically reduces disulfide bonds) before injection [14]. Other disulfide-reducing agents, anionic detergents, proteases, and high or low concentrations of salts also promote the decondensation of nuclear chromatin in mammalian spermatozoa [19, 23, 24, 26–28], again suggesting that the stability of the sperm nucleus is key to the ease with which decondensation occurs. Further evidence for hamster sperm nuclear disulfide bond reduction in the oocyte has been provided by experiments in which decondensation of microinjected sperm nuclei was blocked by treating oocytes with agents that inhibit their ability to reduce disulfide bonds [29]. Conversely, incubation of bovine sperm with DTT has been shown to lead to a partial decondensation of the sperm nucleus [30, 31].

The success of bovine ICSI also depends on proper activation of the oocyte [15, 18, 32]. Mechanical stimulation by the injection pipette alone can only occasionally bring this about in cattle; in more than 95% of cases this stimulus is insufficient [15]. Activation can be induced by a variety of stimuli, including exposure to calcium ionophores [33], ethanol [34, 35], electric currents [33, 36–38], cycloheximide, and 6-dimethylaminopurine (DMAP; a histone kinase inhibitor) [39]. Combining the calcium ionophore, ionomycin, with DMAP has been shown to be particularly effective in inducing bovine oocyte activation, the effects varying with the interval between the sequential treatments with the two compounds [40, 41].

In view of the dependence of sperm nuclear decondensation on the reduction of disulfide bonds, we hypothesized that initiating this process by treating intact sperm before using them for ICSI would facilitate their nuclear decondensation and hence subsequent male PN formation. In this paper, we present evidence that the efficiency of bovine ICSI (judged by the rate of PN formation, the chromosomal constitution of resulting embryos, and their development in vitro) can be improved by sperm pretreatment with DTT. In addition, oocyte activation with ionomycin and DMAP has been shown to be an effective component of the ICSI procedure. However, the viability of the embryos produced in this way with DTT-treated sperm seems limited: although six pregnancies at Day 49 were established by embryo transfer, none was maintained to term.

	MATERIALS AND METHODS

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Experimental Design

The study comprised six experiments. In experiment 1, DTT-treated sperm injection and various stimuli to induce oocyte activation were compared for their effects on cleavage and subsequent development in vitro. The five groups (Table 1) were composed of replicates that were not entirely contemporary, which limited the statistical comparisons that could be made among them. As a consequence of this experiment, the activation of oocytes in all subsequent experiments (experiments 2–6) was accomplished with ionomycin followed 3 h later by DMAP.

In experiments 2–4, ICSI embryos produced with control and DTT-treated spermatozoa were compared for the rates of PN formation 16 h after ICSI (experiment 2), rates of cleavage and development to the blastocyst stage by 192 h (experiment 3), and the ploidy of blastocysts (experiment 4). Replicates comprised control and treated groups on any given day. However, because DTT-treated sperm had to be used before control sperm (to limit their exposure to DTT), and because the efficiency of injection declined with time (through loss of sharpness of the injection pipette), numbers of oocytes in the DTT-treated groups were higher than in the control groups.

In experiment 5, 12 blastocysts produced with DTT-treated sperm were sexed in order to establish whether the presence of a Y chromosome could confirm that they were the product of fertilization rather than parthenogenesis.

Finally, in experiment 6, the viability of blastocysts from Day 8 after ICSI, produced only with DTT-treated sperm, was assessed in vivo; they were transferred to 16 Holstein heifer recipients synchronized (± 1 day) by cloprostenol treatment. The blastocysts were transferred in pairs to the uterine horn ipsilateral to the corpus luteum on Day 7 (n = 12) and Day 8 (n = 4) (estrus = Day 0). Pregnancies were diagnosed by detecting fetal heartbeats by transrectal ultrasonography.

Media

Chemicals were purchased from the Sigma Chemical Company (St. Louis, MO) and media from Gibco (Canadian Life Technologies, Burlington, ON, Canada) unless otherwise specified. The medium used for maturation of cumulus-oocyte-complexes (COCs) was M199 containing Earle's salts, 10% steer serum (Cansera Inc., Rexdale, ON, Canada), 25 mM HEPES, 2.5 mM sodium pyruvate, 1 mM L-glutamine, and 1.0% penicillin-streptomycin (10 000 IU and 10 000 mg/ml, respectively; Pen-Strep; Gibco). The medium used for culture of embryos was M199 containing 2.5 mM sodium pyruvate, 1 mM L-glutamine, and 0.5% penicillin-streptomycin. For both the maturation and the culture media, the pH was adjusted to 7.4 and the osmolality to 280 mOsm/kg. For sperm swim-up, Tyrode's albumin lactate pyruvate medium supplemented with 10 mM HEPES (HEPES-TALP) was used. The micromanipulation medium used for ICSI was Ham's F-10 containing 25 mM HEPES.

Oocyte Preparation

Groups of 15 COCs collected from ovaries harvested at a local slaughterhouse were matured in 50-µl droplets of maturation medium under silicone oil (Dow Corning Medical Fluid 200; Dow Corning, Midland, MI) at 39°C in a humidified atmosphere of 5% CO₂ in air. After 22–24 h in vitro maturation (IVM), the expanded cumulus cells were removed by vortexing for 2 min in Ham's F-10 medium containing 0.5 mg/ml hyaluronidase. Oocytes with an extruded first polar body were subsequently centrifuged in Ham's F-10 for 5 min at 6000 x g to clarify the otherwise dense cytoplasm, and then transferred to drops of preincubated Ham's F-10 medium under silicone oil for sperm injection.

Sperm Preparation

Straws of frozen bull sperm were thawed by immersion in a 37°C water bath for 15 sec. A 0.1-ml aliquot of the thawed semen was layered under 1 ml HEPES-TALP in small culture tubes to permit the sperm to "swim up". To prepare DTT-treated sperm, the HEPES-TALP was supplemented with 5 mM DTT. One hour later, the sperm that had swum up into the top 0.3 ml in each culture tube were collected and washed twice by centrifugation for 10 min at 350 x g in 10 ml HEPES-TALP to remove DTT. In preparation for ICSI, 2 µl of the sperm suspension was transferred to 10 µl of 10% polyvinylpyrrolidone (PVP; M_r 360 000) solution in Ham's F-10 medium under silicone oil. This prevented the spermatozoa from sticking to the inner surface of the micropipette and also reduced their motility.

The morphology of DTT-treated sperm was assessed by phase-contrast microscopy. To assess their viability, DTT-treated sperm were stained with a vital stain (FertiLight; Molecular Probes Inc., Eugene, OR). Sperm stained green (live) and red (dead) were counted on a Zeiss fluorescence microscope (Carl Zeiss, Inc., Thornwood, NJ) fitted with an excitation filter of 365 nm and a barrier filter of 397 nm.

Sperm Microinjection

ICSI was performed at x200 magnification in 30-µl droplets of Ham's F-10 under silicone oil in 60 x 15-mm tissue culture dishes (Falcon; Fisher Scientific, Atlanta, GA) maintained at 37°C on the heated stage of a Leitz inverted microscope (Leitz Wetzlar, Wetzlar, Germany). The injection pipette with an inner diameter at the tip of 8 µm was connected to a pair of Leitz micromanipulators, and the holding pipette was connected to a 1-ml tuberculin syringe. Each spermatozoon for injection was selected from among those in the 10-µl droplet of 10% PVP which stuck by their heads to the bottom of the Petri dish. In the case of DTT-treated sperm, altered morphology of the head (see Results) was used as an additional selection criterion. A selected spermatozoon was immobilized and aspirated tail-first into the injection pipette, which was then moved to the drop containing the centrifuged oocytes to be injected. For injection, the injection pipette was first used to indent the oolemma across approximately five-sixths of the diameter of the oocyte. There, aspiration was used to break the oolemma, give access to the cleared area of the ooplasm, and draw a small volume of ooplasm into the pipette. The spermatozoon and the aspirated ooplasm were then expelled into the oocyte with a minimal volume of PVP solution. About 10 oocytes were placed in the micromanipulation drop to avoid exposure to the Ham's F-10 medium for longer than 20 min. One hour after injection, the oocytes were re-examined, and any from which the spermatozoon could be seen to have been expelled into the perivitelline space were removed.

Activation and Further Development of the Oocytes

In the first experiment (see Experimental Design), four potential means of oocyte activation were compared: injection (with or without introduction of sperm) with no further stimulus, ionomycin alone, or ionomycin plus DMAP by a slight modification of one of the methods originally reported by Susko-Parrish et al. [40]. In brief, the last method consisted of exposing the oocytes to 5 µM ionomycin in HEPES-TALP containing 1 mg/ml BSA (fraction V) for 5 min after injection and then to HEPES-TALP containing 30 mg/ml BSA for 6 min to stop the activation process. Activated oocytes were washed in HEPES-TALP containing 1 mg/ml BSA, cultured in M199 for 3 h to permit extrusion of their second polar body, then transferred to a drop of 1.9 mM DMAP in M199 for 3 h [41]. Activated ICSI oocytes were cocultured with bovine oviductal epithelial cells (BOEC) in sets of 15 in 50-µl drops of M199 as described by Rieger et al. [42]. These cocultures were maintained for 8 days. At Days 2 and 5 (Day 0 = day of injection), the cultures were "fed" by adding 25 µl of serum-free M199 to each drop. Embryonic development was assessed with an inverted microscope at 24-h intervals for up to 192 h after injection.

Cytological Procedures and Embryo Sexing

At 16 h after ICSI, some oocytes (see Results) were fixed overnight in methanol:acetic acid (3:1, v:v) and then stained with 1% aceto-lacmoid to reveal the presence of pronuclei.

At the blastocyst stage, some embryos were prepared and examined for their cytogenetic composition as described by King et al. [43]. Embryos were classified as being mixoploid (embryos with blastomeres of different ploidies), polyploid ( 3n), or diploid (2n).

In order to determine cell numbers, blastocysts at 192 h after ICSI with DTT-treated sperm were fixed in methanol:acetic acid (3:1, v:v) overnight and then stained with 4% Giemsa solution for 10 min. Nuclei were counted at x100 magnification.

The embryos were sexed according to the polymerase chain reaction (PCR) technique of Bredbacka and Peippo [44] and Bredbacka and Bredbacka [45].

Statistical Analysis

In experiments 1 and 3, differences among treatments were analyzed using one-way ANOVA after arc-sine transformation of the proportional data of cleavage and development. Comparison of means among treatments was performed using the Tukey-Kramer multiple comparisons test. In experiments 2 and 4, differences among treatments were analyzed using Yates' corrected chi-square. Differences were considered significant when p < 0.05.

	RESULTS

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General Observations

At the time of cumulus cell removal, approximately 70% of the oocytes matured in vitro had a visible first polar body and uniformly dense ooplasm, which were considered to make them suitable for ICSI. Out of a total of 761 injected oocytes, 57 (8%) were mechanically damaged by the micromanipulation procedure and degenerated within 3–24 h. A further 32 (4%) expelled the injected spermatozoon into the perivitelline space. The damaged oocytes, as well as those with the spermatozoon expelled into the perivitelline space, were not considered to have been successfully injected, leaving 672 of 761 for further consideration.

In preliminary experiments (data not shown), it was found difficult to be sure that the sperm had been successfully injected into noncentrifuged oocytes because of the opacity of the ooplasm. However, when the oocytes were centrifuged, the injected sperm could be easily seen in the ooplasm at x200 magnification. Within approximately 10 h of centrifugation, the lipid had become redistributed throughout the ooplasm. Aspiration of the oolemma to rupture it and gain access to the ooplasm for sperm injection was found to be very efficient. The size (inner diameter 8 µm) of the injection pipette, as well as the spike on its tip, were also found to be critical to the ease and success of bovine ICSI.

It was found in parallel studies [41] that most oocytes (89%, 41 of 46) treated with DMAP directly after activation with ionomycin failed to extrude the second polar body. However, when the activated oocytes were cultured for 3 h before DMAP treatment, the second polar body was extruded in 93% (39 of 42) of them.

After incubating the sperm with 5 mM DTT for 1 h, most (83%, 578 of 696, 6 replicates) still possessed an intact membrane as judged by vital staining 2 h after treatment. However, 46% of treated sperm (163 of 355, 3 replicates) displayed altered morphology; under phase contrast, the heads of these sperm exhibited pronounced bending near the equatorial segment (Fig. 1A), and the midpiece of the tail appeared to be swollen and bent. No such changes were seen in untreated sperm (0 of 227, 2 replicates). Further incubation of the sperm in HEPES-TALP for up to 5 h led to progressive time-dependent decondensation of the nucleus (Fig. 1, B and C). Again, no such decondensation was seen in the untreated sperm (208 examined in 2 replicates).

View larger version (55K): [in this window] [in a new window]   FIG. 1. Morphological changes observed in bovine spermatozoa treated with 5 mM DTT in HEPES-TALP for 1 h, then washed and incubated in HEPES-TALP for 1 h (A), 3 h (B), or 5 h (C). A) Spermatozoa folded near their equatorial segments as seen frontally (a) or from the side (b). B) Spermatozoon with a partially decondensed nucleus. C) Spermatozoon with a fully decondensed nucleus. (Phase contrast; bars = 5 µm).

Experiment 1: Cleavage and Development In Vitro after Various Activation Procedures

As shown in Table 1, cleavage was rare in the sham-injected oocytes of group 1, and only one of them developed to the morula stage. In group 2, the cleavage rate remained low and no further development occurred. In group 3, the rates of cleavage and of development to the blastocyst stage were 17% and 4%, respectively. With the final activation procedure (ionomycin and DMAP), developmental rates were appreciable even without sperm injection (group 4; 42% cleaved, and 11% developed to the blastocyst stage). However, the rates were further amplified in DTT-treated sperm-injected oocytes that were similarly activated (group 5; 61% cleaved, and 24% became blastocysts). The number of nuclei per blastocyst at 192 h after ICSI in group 5 ranged from 69 to 152 (n = 24; mean ± SEM = 121 ± 5).

Experiment 2: Cytological Observations 16 h after ICSI with Control and DTT-Treated Sperm

Oocytes with one PN (± a second polar body) were considered to have been activated whereas those with two PN and a second polar body were considered to have been fertilized. Oocytes were considered to have been unsuccessfully injected if no sperm could be detected. Table 2 shows that the rates of activation, based on these criteria, were 55 of 79 (70%) and 68 of 86 (79%) with the control and DTT-treated sperm, respectively. These rates did not differ significantly. In contrast, Table 2 also shows that the proportion of activated, successfully injected oocytes that had been fertilized was higher 16 h after ICSI with DTT-treated than with control sperm (34 of 57: 60% vs. 9 of 39: 23%; p < 0.001). In both groups, some of the oocytes that contained intact sperm heads remained at metaphase II. The proportion of oocytes in which neither a male PN nor an intact sperm head could be detected were 28 of 79 (35%) and 17 of 86 (20%) in the control and DTT-treated groups, respectively, and did not differ significantly.

Experiment 3: In Vitro Development after ICSI with Control and DTT-Treated Sperm

The rates of cleavage obtained after ICSI with control and DTT-treated sperm were not significantly different (38 of 61: 62%, and 64 of 93: 69%, respectively). However, the proportion of injected oocytes that developed into blastocysts was significantly higher in the treated than in the control group (22 of 93: 24% vs. 6 of 61: 10%; p < 0.005).

Experiment 4: Chromosomal Analysis of Blastocysts Produced by ICSI with Control and DTT-Treated Sperm

Sixty-eight blastocysts in the two groups were used to determine their ploidy (Table 3). There was no significant difference in the frequency of abnormalities between those produced with control and with DTT-treated sperm (29% and 24% mixoploid and polyploid blastocysts, respectively, in the control group and 22% and 18%, respectively, in the DTT-treated group). The rates of diploid blastocyst formation in control and DTT-treated groups were not significantly different (47% and 61%, respectively).

Experiment 5: The Sexing of Blastocysts Produced by ICSI with DTT-Treated Sperm

Out of 12 blastocyst-stage embryos produced with DTT-treated sperm, 4 were diagnosed as males and 7 as females, and in one a definite diagnosis could not be made by PCR.

Experiment 6: Transfers of Blastocysts Produced by ICSI with DTT-Treated Sperm

Out of 16 recipient heifers that each received 2 ICSI embryos, 6 (38%) were confirmed to be pregnant by detecting fetal heart beats by transrectal ultrasonography at Day 49 of gestation. However, none of these pregnancies were carried to term.

	DISCUSSION

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This study has demonstrated that the efficiency of ICSI in cattle can be improved by modifying the usual ways of preparing both the spermatozoon and the oocyte for the procedure.

Treating the sperm with DTT before using them for ICSI clearly increased the rate at which they gave rise to male pronuclei in injected oocytes. This result corroborates a previous report [14] that DTT-treated bovine sperm nuclei, isolated by sonication, were able to participate more fully in DNA synthesis in the hamster oocyte due to more rapid decondensation and PN formation. This improvement may be caused by the reduction of stabilizing disulfide bonds. As it passes through the epididymis, the mammalian sperm nucleus becomes very resistant to physical and chemical disruption due to the conversion of protamine sulfhydryl groups to disulfides [19–21]. Bovine sperm nuclei have particularly strong disulfide bonds [14], which could be expected to result in hyperstabilized chromatin, preventing the sperm nucleus from decondensing [46].

In the present investigation, in contrast to previous studies of the isolated sperm nucleus, intact spermatozoa were treated with DTT. After incubation of the sperm with DTT for 1 h, the membrane of most sperm remained intact as judged by vital staining 2 h after treatment. However, phase-contrast microscopy showed that the morphology of the sperm head and midpiece of the tail was frequently altered. The heads of these sperm could be seen to be folded back near the equatorial segment, and the midpiece of the tail appeared swollen or bent. Further incubation, for up to 5 h after treatment in HEPES-TALP, led to partial or complete decondensation of the sperm nucleus (Fig. 1). These morphological changes suggest that the sperm membrane, particularly near the equatorial segment and the midpiece, may also become weakened by the DTT treatment, which may also help to decondense the sperm nucleus following ICSI. Others have suggested that DTT treatment caused morphological changes in perforatorium, midpiece, and head in mammalian sperm [19, 30, 31, 47].

Our study has also demonstrated that an oocyte activation procedure using ionomycin and DMAP [41], developed in parallel with the present work, is more efficient than is ionomycin alone. It is known that calcium plays an important part in the intracellular signaling responsible for the initiation and propagation of oocyte activation in all mammalian species. During ICSI, puncture of the oolemma [6] or aspiration of cytoplasm [48] of human oocytes with a micropipette can contribute to oocyte activation. Rabbit oocytes can also be activated by pricking the plasma membrane with a micropipette, but oocytes treated with 10 µM calcium ionophore for 10 min after ICSI develop at a higher rate [49]. Bovine oocytes are not sufficiently activated by the injection procedure itself to complete meiosis [15, 17, 32]. Keefer et al. [15] demonstrated that oocyte activation by calcium ionophore A23187 was required in order to get male PN formation and cleavage following sperm injection. Using this procedure, Goto [17] achieved rates of 15% cleavage (84 of 557) and 7% development into blastocysts (12 of 165). Tocharus et al. [50] also reported that when oocytes were activated with 7% ethanol for 5 min before ICSI, development rates to the 2-cell and blastocyst stages were 68% (74 of 108) and 16% (17 of 108), respectively.

In choosing an activation procedure for the present study, a preliminary study (not presented) showed that when oocytes were stimulated electrically (DC 1.5 kv/cm, 60 µsec, twice, in 0.3 M mannitol solution containing 100 µM CaCl₂ and MgCl₂) using a BTX electrocell manipulator (BTX, San Diego, CA) or with 7% ethanol after ICSI, cleavage rates (4 of 30: 13%, and 3 of 41: 7%, respectively) and rates of development into blastocysts (1 of 30: 3%, and 1 of 41: 2%, respectively) were very low. Therefore, for oocyte activation, we decided to use one of the ionomycin and DMAP methods of Susko-Parrish et al. [40] which seemed appropriate to ICSI; the regimen with a 3-h interval between ionomycin activation and DMAP treatment allowed time for the extrusion of the second polar body. In our experience [41], the proportion of oocytes extruding the second polar body after this treatment can be much higher than was reported previously by Susko-Parrish et al. (93% vs. ~33%). Perhaps different intensities of selection of oocytes used for activation could account for this discrepancy.

Although a satisfactory rate of oocyte activation was achieved using this method, often it resulted in abnormal embryos. Of the 51 embryos in the DTT-treated group subjected to chromosome analysis, 31 (61%) were diploid, while the others (39%) were mixoploid or polyploid. This high proportion of mixoploidy and polyploidy contrasts markedly with the 17% of chromosomal abnormalities estimated to occur in bovine embryos resulting from conventional in vitro fertilization [51]. Mixoploidy or polyploidy may be due to the failure of second polar body extrusion or, alternatively, to parthenogenesis. The latter explanation seems the more likely, given the high rate of second polar body extrusion obtained with the activation method used. Because the activation procedure itself, without sperm injection, gave rise to blastocysts from about 11% of treated oocytes, it was important to confirm that the improved rate of blastocyst formation (24%) after ICSI was not due simply to an increased rate of parthenogenesis. The demonstration of the Y chromosome in one-third (4 of 12) of ICSI embryos sexed by PCR indicates that those embryos, at least, had been fertilized by the injected sperm.

Centrifugation of the oocytes after removal of the cumulus cells following IVM was also felt to facilitate ICSI considerably, although direct comparison with noncentrifuged controls was not undertaken. Centrifugation clarified the ooplasm by polarizing the lipid, which is well known to obscure cellular organelles in the oocytes of many ungulates. Wall et al. [52] first used this procedure to facilitate pronuclear injection in order to produce transgenic cattle and pig zygotes, and it has since been used to subject pig zygotes to "lipidectomy" before their cryopreservation [53] and enucleation of bovine oocytes for use in nuclear transfer [54]. Tatham et al. [55] have recently suggested that centrifugation (at 15 800 x g for 10 min) of oocytes before ICSI would aid in better visualization of the injected spermatozoon. In the present study, we found that centrifugation of the oocyte at 6000 x g for 5 min made it possible to see the spermatozoon as it was expelled from the injection pipette into the ooplasm. As a result, the injection volume could be reduced to a minimum. Centrifugation at this force is less rigorous than the procedures used by Wall et al. [52] on bovine and pig zygotes (15 000 x g for 3 min) or by Nagashima et al. [53] on pig zygotes (12 000 x g for 8 min). Parallel studies (not presented) showed that centrifugation did not damage the oocytes; they could be fertilized in vitro at the same rate as noncentrifuged oocytes. Others have also shown that centrifugation does not impede subsequent development [56]. This may be because the polarization of the ooplasm is only temporary; in the present study, the lipid became redistributed throughout the ooplasm by about 10 h after centrifugation.

Although only a small number of embryo transfers were performed, 6 out of 16 (38%) recipient heifers were confirmed to be pregnant at Day 49. Furthermore, the detection of fetal heartbeats suggests that the conceptuses resulted from fertilization rather than parthenogenesis because heartbeats were not found in "fetal masses" detected in utero after the transfer of bovine parthenotes [40, 57]. However, none of the pregnancies proceeded to term, suggesting that the viability of the blastocysts produced by ICSI was limited.

In summary, the present data indicate that the pretreatment of sperm with DTT and the activation of oocytes with ionomycin and DMAP are useful for facilitating sperm decondensation and subsequent PN formation after ICSI. It is also suggested that oocyte centrifugation before injection is beneficial to the procedure. Further studies, particularly of oocyte activation, are clearly needed to develop an ICSI technique that will regularly result in chromosomally normal embryos, fetuses, and calves.

	ACKNOWLEDGMENTS

 
The authors thank Drs. W.A. King and C.L. Keefer for critical reading of this manuscript, Mrs. Suzanne Manning and Ms. Léanne Bradley for technical support, and Dr. Ping Xia for her help in fabrication of injection pipettes.

	FOOTNOTES

 
¹ Supported by Natural Sciences and Engineering Research Council (NSERC) of Canada and the Ontario Ministry of Agriculture, Food and Rural Affairs (OMAFRA).

² Correspondence. FAX: 519 824 1643; kbetter{at}uoguelph.ca

Accepted: June 4, 1998.

Received: October 29, 1997.

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