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In vitro maturation of horse oocytes
Theriogenology
42:345-349,1994
IN VITRO MATURATION OF HORSE OOCYTES H. Aim and H. Tomer Department of Reproductive Biology Research Institute for Biology of Farm Animals Dummerstorf, Germany Received for publication: Accepted:
May 25, 1993 June 14, 1994
ABSTRACT Ovaries collected from slaughtered mares of unknown reproductive history were transported to the laboratory, and their oocytes were recovered and cultured in modified TCM 199 supplemented with 20 % horse serum and additional granulosa cells. To characterize the ovaries, the size and number of follicles were counted. To determine the time required for nuclear maturation, oocytes were fixed either after 18 h (n=23), 24 h (n=50), or 30 h (n=33) of culture. After co-culture with granulosa cells most oocytes reached metaphase II (M II) by 30 h (72.7 %), After 24 h of maturation only 56.0 % of the cultured oocytes had reached metaphase II (M 11). Key words: mare, oocytes, collection, granulosa cells, maturation in vitro INTRODUCTION Maturation and fertilization in vitro has now been used successfully in the human, cow, pig, rhesus monkey and several laboratory animal species. In contrast to the situation of many domestic animals, development of IVM/IVF procedures has been slow in equids. This has been partly due to the limited availability of equine oocytes, but also to the difficulty in capacitating stallion spermatozoa outside the mare’s reproductive tract (3,17). The time required for maturation of equine oocytes in vitro has been reported, but the results are conflicting. Fulka and Okolski (6) used modified TCM 199 and found that the time required for nuclear maturation is more than 24 h but less than 40 h. The results of Willis et al. (15) indicate that the culture of equine oocytes in vitro for 15 and 32 h resulted in significant (P
The aim of the present study was to define the number and size of follicles and to describe a method for the recovery and maturation of horse oocytes.
Copyright
0 1994 Butterworth-Heinemann
Theriogenology
346 MATERIALS AND METHODS
Ovaries were obtained from 23 mares of unknown reproductive history at an abottoir. Immediately after slaugther, the ovaries were transported in a thermos container in TCM 199 to the laboratory within 2.5 to 4 h. The container and the medium were prewarmed and the transport was carried out at 20-22“ C. Upon reaching the laboratory the ovaries were washed in fresh medium and measured. After removing the tunica albuginea the visible follicles on the surface of the ovaries were measured, and their number and size were determined. Oocytes were recovered either by aspiration through IS-gauge needles or by rupturing of isolated follicles. Each isolated follicle was put in a petri dish with fresh medium, and the follicles were opened and pressed with tweezers and a needle under the stereo microscope in order to observe the release of the oocyte. All preparations were done in a cell culture laboratory at room temperature. The oocytes were collected in TCM 199 + 10 % horse serum and held at room temperature until all oocytes had been collected. The oocytes were then washed twice, and only oocytes with complete, compact multilayered cumulus and dark evenly granulated cytoplasm were used for maturation in vitro. The cumulus-oocyte-complexes (COC) were cultured at 38.5 “C under 5 % CO2 in 100% humidified air in maturation medium TCM 199 (Sigma Chemical Co., Deisenhofen, Germany) containing 20 % (v/v) heat treated horse serum (Sigma H-l 138), 100 IU penicillin/ml (Sigma), and 100 ug strepromycin/ml (Sigma). To simulate the natural environment and on the basis of the results in bovine oocyte maturation (1) mare granulosa cells (2 to 5xlOWml) were added to the medium. The granulosa cell preparation was carried out by the methods previously described by Lu et al. (9): granulosa cells were collected from the dissection medium in which the COC were dissected from the follicles and centrifuged twice for 5 min at 500 g. The final pellet of granulosa cells was suspended in the maturation medium, and the resulting suspension was passed through an IS-gauge needle attached to a l-ml syringe, in order to disperse the cells. The co-culture of oocytes and granulosa cells was carried out in petri dishes in 2 ml of medium in a flux system designed to provide gentle agitation but not to achieve cellular confluence. At the end of culture, the cumulus cells were removed by treatment with sodium citrate (3 %). To classify the stage of nulear maturation as germinal vesicle (GV), germinal vesicle breakdown (GVBD), meta- and anaphase I (M I, A I) or metaphase II (M II) oocytes were fixed with acetic alcohol and stained with 1 % aceto-orcein. W-square significant.
analysis was used, and all results with P
statistically
RESULTS The mean number of visible follicles per ovary (n = 40 ovaries) was 9.4 in total. After removing the tunica albuginea the visible follicles on the surface of the ovaries were measured. The following are the number and size that were determined: 6.1 of the follicles were smaller than 15 mm, 3.0 were between 15 and 30 mm and 0.3 were larger than 30 mm in diameter. Altogether, 365 follicles were either aspirated or isolated (Table 1). The recovery rate overall was 40.3 %. Independent of the follicle size, the rate of oocytes recovered by isolation was
347
lherjogenoiogy
signikantly higher than by aspiration (50.0 vs 31.2 %). The difference in oocyte recovery between aspiration and isolation increased when the follicles were classified according to their diameters. In the follicles Cl5 mm in diameter 61.5 % of the oocytes were recovered by isolation and only 32.3 % by aspiration. However, no significant difference in the rate of recovery was found in foilicles 15 to 30 mm in diameter. Both methods of oocyte recovery yielded similar results (29.1 vs 3 1.5 %). There was no difference in tbe number of recovered compact cumulus-ooc~e-com~Iexes from different-size follicles, and there was no significant difference in the number of compact cumulusoocyte-complexes between follicles smaller than 15 mm (42.9 %) and follicles larger than 15 mm (43.7 %). Table 1. Oocyte recovery in mares by aspiration or isolation based on follicle size Type of recovery
Recovery rate (total, %)
421130 (32.3)a
591189 (31.2)a
Aspiration (n=22 ovaries)
Recovery of oocytes of follicles < 15 mm 15-30 mm > 30 mm 17154 (31.5)
Of5 (0)
88/176 72/l 17 16/55 014 Isolation (0) (50,O)b (61.5)b (29.1) (n=27 ovaries) a,b Difkent superscripts in the same cohunns indicate significant differences between groups (P
The influence of culture time is shown in Table 2. In each experiment, similar proportions of oocytes were fixed after 0,18,24 and 30 h. After 18 h of incubation, the same number of oocytes were in both the resumption of meiosis and in M II (39.1 and 34.8 %, respectively). Less than 10 % were still at the GV stage; 6 h later about 56 % of the oocytes had reached M II, and 20 % of the oocytes were in late meiotic stages (GVBD-A I). The results indicate that culture of equine oocytes in vitro for 30 h increases the number (72.7 Oh)in M II stages. Table 2. Influence of culture tie Time of incubation (hours) 0
18 24 30
on the development of equine oocytes in vitro
No. of oocytes
Immature (Gv) n (%)
Resumption of meiosis (GVBD-A I) n (%)
Mature (metaphase II) n (Oh)
35 23 50 33
17 (48.6) 2 (8.7) 0 (0) o (0)
12 (34.3)a 9 (39.1) 10 (20.0) 0 (0)b
2 (5.7)a 8 (34.g)b 28 (56.0 b 24 (72.7) b S
Degeneration n (%)
4 (11.4) 4 (17.4) 12 (24.0) 9 (27.3)
a,b Different superscripts in the same columns indicate significant differences between groups (P~O.05).
348
Theriogenolog
y
We found that oocytes which did not develop in time degenerated. After 24 h of incubation 11 of 12 oocytes which showed degenerative changes in chromatin configuration were in the GV stage (3 oocytes) or in the other stages of meiosis, up to A I (8 oocytes). Only 1 oocyte at the M II stage degenerated. After 30 h of incubation, all oocytes which did not develop to M II showed degenerated chromatin configuration (8 oocytes), and only 1 oocyte degenerated at the M II stage. DISCUSSION Since the supply of equine ovaries is limited, the number of oocytes per ovary is a critical factor in using in vitro reproductive technology. Aspiration of follicular fluid containing oocytes is the prime method of extracting immature oocytes in different species of animals. Utilizing this method in cattle, 40 to 80 % of the follicular oocytes can be recovered (13). Comparison of the 2 methods of surface cutting and aspiration in sheep concluded that surface cutting is the superior method (7.3 vs 2.1 oocytes/ovary) for the recovery of follicular oocytes (8). In our study, 3 1.2 % of mare follicular oocytes were recovered by aspiration. A lower number of oocytes was obtained by Choi et al. (4) with the aspiration method, but they increased this by using the slicing technique (1.7 vs 4.1 oocytes/ovary). Rupturing isolated follicles proved to be twice as effective as either of the above methods in recovering oocytes from mares. Despite the relatively high recovery rate of oocytes by rupturing (6 1.5 % from follicles < 15 mm), this method is labor intensive and time-consuming. It required about 1 to 1.5 h per ovary to prepare the ovary, isolate the follicle and recover the oocytes. However, since the availability of equine oocytes is limited, rupturing of isolated follicles is the preferred method. In our study, a sufficient number of oocy-tes with an intact cumulus investment (42.9 % from follicles cl5 mm, 43.7 % from follicles 15 to 30 mm) was recovered. These results are similar to those of Choi et al. (4), who recovered about 55 % of oocytes with a compact, circular cumulus, but in contrast with those of Okolski et al. (1 l), who reported that follicular aspiration resulted in the collection of oocytes largely denuded of cumulus cells. The percentage of denuded oocytes in the present study was less than 10 % (5.3 % of follicles cl5 mm, 9.4 % of follicles 15 to 30 mm). The length of time of maturation of horse oocytes appears to be longer than that of most other species. Results of the effects of culture period indicate that culture of equine oocytes in vitro for 30 h resulted in an increase in M II stages (72.7 %), similar to the results of Hinrichs et al. (7) who observed a maturation rate of 75 % after 32 h of incubation and only 42 % after 24 h of incubation. Zhang et al. (17) reported similar results, with 61 % M II after 30 h of culture. The period between the time of ovary recovery and the initiation of oocyte culture can have an effect on development. Periods varied between 3 and 5 h in our experiments. The same interval was used by Okolski et al. (12), for a maturation rate of 37.5 % for M II after 26 h and 53.3 % after 36 h of incubation. After 24 h of incubation, the proportion of matured oocytes in our experiments was even higher (56 % for M II). It seems that co-culture with granulosa cells has a beneficial effect on oocyte nuclear maturation. Supplementation of the maturation medium with additional granulosa cells was also found to be benetltial for achieving full developmental competence of ovine and bovine oocytes (2, 10, 14). Investigation is needed to determine if the fertiliiability of co-cultured equine oocytes can also be improved.
349
Theriogenology
The results of the present study indicate that recovered oocytes with a compact cumulus and a dark, evenly granulated cytoplasm were competent for maturation in vitro and that the period of maturation takes more than 24 h, as in the bovine. The co-culture of oocytes and granulosa cells resulted in a high proportion of M II-oocytes. Our methodology should provide a basis for further investigation and testing of the competence of the recovered oocytes for fertilization in vitro. REFERENCES 1. Alm H, Tomer H, Kanitz W. Development capacity of bovine oocytes in vitro after different FSH/LH ratios in superovulation treatment. 12th Int Cong Anim Reprod 1992; 1:300-302. 2. Alm H, Kauffold P, Makarowa SN,. Sirotkin AV. Der EintluB von Granulosazellen auf die Reifung, Befruchtung und Furchung m vitro. Arch exper Vet med 1990; 4483-91. 3. Blue BJ, McKinnon AO, Squires EL, Seidel JR, Muscari KT. Capacitation of stallion spematozoa and fertilization of equine oocytes in vitro. Equine Vet J 1989; 8 (Suppl): 11 l116. 4. Choi YH, Hochi S,. Braun J, Oguri N. In vitro maturation of equine oocy-tes collected by aspiration and addmonal slicing of ovaries, Theriogenology 1993; 39:200 abstr. 5. Del Camp0 MR, Donoso MX, Parrish JJ. In vitro maturation of equine oocytes. Theriogenology 1992; 37:200 abstr. 6. Fulka J Jr, Okolski A. Culture ofhorse oocytes in vitro. J Reprod Fertil 1981; 61:213-215. 7. Hinrichs K, Friedman PP, Martin MG. Evaluation of in vitro maturation of horse oocytes using fluorescence microscopy. 12th Int Cong Anim Reprod 1992; 1:330-332. 8. Kay GW, Frylinck S. Recovery of ovine follicular oocytes: Effect of different methodes on yield and quality. 12th Int Cong Anim Reprod 1992; 1:345-347. 9. Lu M, Gordon I, Gallagher M, McGovern H. Pregnancy established in cattle by transfer of embryos derived from in vitro fertilization of oocytes matured in vitro. Vet Record 1987; 121:259-260. 10. Mochizuki H, Fukui Y, Ono H. Effect of the number of granulosa cells added to culture medium for in vitro maturation, fertilization and development of bovine oocytes. Theriogenology 1991; 36:973-986. 11. Okolski A, Babusik P, Tischner M, Lietz W. Evaluation of mare oocyte collection methods and stallion sperm penetration of zona-free hamster ova. J Reprod Fertil 1987; 35 (Suppl): 191-196. 12. Okolski A, Bezard J, Magistrini M, Palmer E. Maturation of oocytes from normal and atretic J Reprod Fertil 1991; 44 equine ovarian folhcles as affected by steroid concentration. (Suppl):385-392. 13. Spitschak M, Becker .F, Kanitz W. Ultraschallgestutzte Untersuchungen zum z;zrelwachstum und Folhkelreifung beim Rind. Reprod Dom Anim 1993; 2 (Suppl): 167 14. Staigmiller RB, Moor RM Effect of follicle cells on the maturation and developmental competence of ovine oocytes matured outside the follicle. Gamete Res 1984; 9:221-229. 15. Willis P, Candle AB, Fayrer-Hosken RA. Equine oocyte in vitro maturation: Influence of sera, time and hormones. Molec Reprod Devel 1991; 30:360-368. 16. Willis P, Candle AB, Fayrer-Hosken RA. Ultrastructure of equine oocytes matured in vitro. Theriogenology 1993; 39:340 abstr. 17. Zhang JJ, Boyle MS, Allen WR, Galli C. Recent studies on in vivo fertilization of in vitro matured horse oocytes. Equine Vet J 1989; 8 (Suppl): 101-104.
42:345-349,1994
IN VITRO MATURATION OF HORSE OOCYTES H. Aim and H. Tomer Department of Reproductive Biology Research Institute for Biology of Farm Animals Dummerstorf, Germany Received for publication: Accepted:
May 25, 1993 June 14, 1994
ABSTRACT Ovaries collected from slaughtered mares of unknown reproductive history were transported to the laboratory, and their oocytes were recovered and cultured in modified TCM 199 supplemented with 20 % horse serum and additional granulosa cells. To characterize the ovaries, the size and number of follicles were counted. To determine the time required for nuclear maturation, oocytes were fixed either after 18 h (n=23), 24 h (n=50), or 30 h (n=33) of culture. After co-culture with granulosa cells most oocytes reached metaphase II (M II) by 30 h (72.7 %), After 24 h of maturation only 56.0 % of the cultured oocytes had reached metaphase II (M 11). Key words: mare, oocytes, collection, granulosa cells, maturation in vitro INTRODUCTION Maturation and fertilization in vitro has now been used successfully in the human, cow, pig, rhesus monkey and several laboratory animal species. In contrast to the situation of many domestic animals, development of IVM/IVF procedures has been slow in equids. This has been partly due to the limited availability of equine oocytes, but also to the difficulty in capacitating stallion spermatozoa outside the mare’s reproductive tract (3,17). The time required for maturation of equine oocytes in vitro has been reported, but the results are conflicting. Fulka and Okolski (6) used modified TCM 199 and found that the time required for nuclear maturation is more than 24 h but less than 40 h. The results of Willis et al. (15) indicate that the culture of equine oocytes in vitro for 15 and 32 h resulted in significant (P
The aim of the present study was to define the number and size of follicles and to describe a method for the recovery and maturation of horse oocytes.
Copyright
0 1994 Butterworth-Heinemann
Theriogenology
346 MATERIALS AND METHODS
Ovaries were obtained from 23 mares of unknown reproductive history at an abottoir. Immediately after slaugther, the ovaries were transported in a thermos container in TCM 199 to the laboratory within 2.5 to 4 h. The container and the medium were prewarmed and the transport was carried out at 20-22“ C. Upon reaching the laboratory the ovaries were washed in fresh medium and measured. After removing the tunica albuginea the visible follicles on the surface of the ovaries were measured, and their number and size were determined. Oocytes were recovered either by aspiration through IS-gauge needles or by rupturing of isolated follicles. Each isolated follicle was put in a petri dish with fresh medium, and the follicles were opened and pressed with tweezers and a needle under the stereo microscope in order to observe the release of the oocyte. All preparations were done in a cell culture laboratory at room temperature. The oocytes were collected in TCM 199 + 10 % horse serum and held at room temperature until all oocytes had been collected. The oocytes were then washed twice, and only oocytes with complete, compact multilayered cumulus and dark evenly granulated cytoplasm were used for maturation in vitro. The cumulus-oocyte-complexes (COC) were cultured at 38.5 “C under 5 % CO2 in 100% humidified air in maturation medium TCM 199 (Sigma Chemical Co., Deisenhofen, Germany) containing 20 % (v/v) heat treated horse serum (Sigma H-l 138), 100 IU penicillin/ml (Sigma), and 100 ug strepromycin/ml (Sigma). To simulate the natural environment and on the basis of the results in bovine oocyte maturation (1) mare granulosa cells (2 to 5xlOWml) were added to the medium. The granulosa cell preparation was carried out by the methods previously described by Lu et al. (9): granulosa cells were collected from the dissection medium in which the COC were dissected from the follicles and centrifuged twice for 5 min at 500 g. The final pellet of granulosa cells was suspended in the maturation medium, and the resulting suspension was passed through an IS-gauge needle attached to a l-ml syringe, in order to disperse the cells. The co-culture of oocytes and granulosa cells was carried out in petri dishes in 2 ml of medium in a flux system designed to provide gentle agitation but not to achieve cellular confluence. At the end of culture, the cumulus cells were removed by treatment with sodium citrate (3 %). To classify the stage of nulear maturation as germinal vesicle (GV), germinal vesicle breakdown (GVBD), meta- and anaphase I (M I, A I) or metaphase II (M II) oocytes were fixed with acetic alcohol and stained with 1 % aceto-orcein. W-square significant.
analysis was used, and all results with P
statistically
RESULTS The mean number of visible follicles per ovary (n = 40 ovaries) was 9.4 in total. After removing the tunica albuginea the visible follicles on the surface of the ovaries were measured. The following are the number and size that were determined: 6.1 of the follicles were smaller than 15 mm, 3.0 were between 15 and 30 mm and 0.3 were larger than 30 mm in diameter. Altogether, 365 follicles were either aspirated or isolated (Table 1). The recovery rate overall was 40.3 %. Independent of the follicle size, the rate of oocytes recovered by isolation was
347
lherjogenoiogy
signikantly higher than by aspiration (50.0 vs 31.2 %). The difference in oocyte recovery between aspiration and isolation increased when the follicles were classified according to their diameters. In the follicles Cl5 mm in diameter 61.5 % of the oocytes were recovered by isolation and only 32.3 % by aspiration. However, no significant difference in the rate of recovery was found in foilicles 15 to 30 mm in diameter. Both methods of oocyte recovery yielded similar results (29.1 vs 3 1.5 %). There was no difference in tbe number of recovered compact cumulus-ooc~e-com~Iexes from different-size follicles, and there was no significant difference in the number of compact cumulusoocyte-complexes between follicles smaller than 15 mm (42.9 %) and follicles larger than 15 mm (43.7 %). Table 1. Oocyte recovery in mares by aspiration or isolation based on follicle size Type of recovery
Recovery rate (total, %)
421130 (32.3)a
591189 (31.2)a
Aspiration (n=22 ovaries)
Recovery of oocytes of follicles < 15 mm 15-30 mm > 30 mm 17154 (31.5)
Of5 (0)
88/176 72/l 17 16/55 014 Isolation (0) (50,O)b (61.5)b (29.1) (n=27 ovaries) a,b Difkent superscripts in the same cohunns indicate significant differences between groups (P
The influence of culture time is shown in Table 2. In each experiment, similar proportions of oocytes were fixed after 0,18,24 and 30 h. After 18 h of incubation, the same number of oocytes were in both the resumption of meiosis and in M II (39.1 and 34.8 %, respectively). Less than 10 % were still at the GV stage; 6 h later about 56 % of the oocytes had reached M II, and 20 % of the oocytes were in late meiotic stages (GVBD-A I). The results indicate that culture of equine oocytes in vitro for 30 h increases the number (72.7 Oh)in M II stages. Table 2. Influence of culture tie Time of incubation (hours) 0
18 24 30
on the development of equine oocytes in vitro
No. of oocytes
Immature (Gv) n (%)
Resumption of meiosis (GVBD-A I) n (%)
Mature (metaphase II) n (Oh)
35 23 50 33
17 (48.6) 2 (8.7) 0 (0) o (0)
12 (34.3)a 9 (39.1) 10 (20.0) 0 (0)b
2 (5.7)a 8 (34.g)b 28 (56.0 b 24 (72.7) b S
Degeneration n (%)
4 (11.4) 4 (17.4) 12 (24.0) 9 (27.3)
a,b Different superscripts in the same columns indicate significant differences between groups (P~O.05).
348
Theriogenolog
y
We found that oocytes which did not develop in time degenerated. After 24 h of incubation 11 of 12 oocytes which showed degenerative changes in chromatin configuration were in the GV stage (3 oocytes) or in the other stages of meiosis, up to A I (8 oocytes). Only 1 oocyte at the M II stage degenerated. After 30 h of incubation, all oocytes which did not develop to M II showed degenerated chromatin configuration (8 oocytes), and only 1 oocyte degenerated at the M II stage. DISCUSSION Since the supply of equine ovaries is limited, the number of oocytes per ovary is a critical factor in using in vitro reproductive technology. Aspiration of follicular fluid containing oocytes is the prime method of extracting immature oocytes in different species of animals. Utilizing this method in cattle, 40 to 80 % of the follicular oocytes can be recovered (13). Comparison of the 2 methods of surface cutting and aspiration in sheep concluded that surface cutting is the superior method (7.3 vs 2.1 oocytes/ovary) for the recovery of follicular oocytes (8). In our study, 3 1.2 % of mare follicular oocytes were recovered by aspiration. A lower number of oocytes was obtained by Choi et al. (4) with the aspiration method, but they increased this by using the slicing technique (1.7 vs 4.1 oocytes/ovary). Rupturing isolated follicles proved to be twice as effective as either of the above methods in recovering oocytes from mares. Despite the relatively high recovery rate of oocytes by rupturing (6 1.5 % from follicles < 15 mm), this method is labor intensive and time-consuming. It required about 1 to 1.5 h per ovary to prepare the ovary, isolate the follicle and recover the oocytes. However, since the availability of equine oocytes is limited, rupturing of isolated follicles is the preferred method. In our study, a sufficient number of oocy-tes with an intact cumulus investment (42.9 % from follicles cl5 mm, 43.7 % from follicles 15 to 30 mm) was recovered. These results are similar to those of Choi et al. (4), who recovered about 55 % of oocytes with a compact, circular cumulus, but in contrast with those of Okolski et al. (1 l), who reported that follicular aspiration resulted in the collection of oocytes largely denuded of cumulus cells. The percentage of denuded oocytes in the present study was less than 10 % (5.3 % of follicles cl5 mm, 9.4 % of follicles 15 to 30 mm). The length of time of maturation of horse oocytes appears to be longer than that of most other species. Results of the effects of culture period indicate that culture of equine oocytes in vitro for 30 h resulted in an increase in M II stages (72.7 %), similar to the results of Hinrichs et al. (7) who observed a maturation rate of 75 % after 32 h of incubation and only 42 % after 24 h of incubation. Zhang et al. (17) reported similar results, with 61 % M II after 30 h of culture. The period between the time of ovary recovery and the initiation of oocyte culture can have an effect on development. Periods varied between 3 and 5 h in our experiments. The same interval was used by Okolski et al. (12), for a maturation rate of 37.5 % for M II after 26 h and 53.3 % after 36 h of incubation. After 24 h of incubation, the proportion of matured oocytes in our experiments was even higher (56 % for M II). It seems that co-culture with granulosa cells has a beneficial effect on oocyte nuclear maturation. Supplementation of the maturation medium with additional granulosa cells was also found to be benetltial for achieving full developmental competence of ovine and bovine oocytes (2, 10, 14). Investigation is needed to determine if the fertiliiability of co-cultured equine oocytes can also be improved.
349
Theriogenology
The results of the present study indicate that recovered oocytes with a compact cumulus and a dark, evenly granulated cytoplasm were competent for maturation in vitro and that the period of maturation takes more than 24 h, as in the bovine. The co-culture of oocytes and granulosa cells resulted in a high proportion of M II-oocytes. Our methodology should provide a basis for further investigation and testing of the competence of the recovered oocytes for fertilization in vitro. REFERENCES 1. Alm H, Tomer H, Kanitz W. Development capacity of bovine oocytes in vitro after different FSH/LH ratios in superovulation treatment. 12th Int Cong Anim Reprod 1992; 1:300-302. 2. Alm H, Kauffold P, Makarowa SN,. Sirotkin AV. Der EintluB von Granulosazellen auf die Reifung, Befruchtung und Furchung m vitro. Arch exper Vet med 1990; 4483-91. 3. Blue BJ, McKinnon AO, Squires EL, Seidel JR, Muscari KT. Capacitation of stallion spematozoa and fertilization of equine oocytes in vitro. Equine Vet J 1989; 8 (Suppl): 11 l116. 4. Choi YH, Hochi S,. Braun J, Oguri N. In vitro maturation of equine oocy-tes collected by aspiration and addmonal slicing of ovaries, Theriogenology 1993; 39:200 abstr. 5. Del Camp0 MR, Donoso MX, Parrish JJ. In vitro maturation of equine oocytes. Theriogenology 1992; 37:200 abstr. 6. Fulka J Jr, Okolski A. Culture ofhorse oocytes in vitro. J Reprod Fertil 1981; 61:213-215. 7. Hinrichs K, Friedman PP, Martin MG. Evaluation of in vitro maturation of horse oocytes using fluorescence microscopy. 12th Int Cong Anim Reprod 1992; 1:330-332. 8. Kay GW, Frylinck S. Recovery of ovine follicular oocytes: Effect of different methodes on yield and quality. 12th Int Cong Anim Reprod 1992; 1:345-347. 9. Lu M, Gordon I, Gallagher M, McGovern H. Pregnancy established in cattle by transfer of embryos derived from in vitro fertilization of oocytes matured in vitro. Vet Record 1987; 121:259-260. 10. Mochizuki H, Fukui Y, Ono H. Effect of the number of granulosa cells added to culture medium for in vitro maturation, fertilization and development of bovine oocytes. Theriogenology 1991; 36:973-986. 11. Okolski A, Babusik P, Tischner M, Lietz W. Evaluation of mare oocyte collection methods and stallion sperm penetration of zona-free hamster ova. J Reprod Fertil 1987; 35 (Suppl): 191-196. 12. Okolski A, Bezard J, Magistrini M, Palmer E. Maturation of oocytes from normal and atretic J Reprod Fertil 1991; 44 equine ovarian folhcles as affected by steroid concentration. (Suppl):385-392. 13. Spitschak M, Becker .F, Kanitz W. Ultraschallgestutzte Untersuchungen zum z;zrelwachstum und Folhkelreifung beim Rind. Reprod Dom Anim 1993; 2 (Suppl): 167 14. Staigmiller RB, Moor RM Effect of follicle cells on the maturation and developmental competence of ovine oocytes matured outside the follicle. Gamete Res 1984; 9:221-229. 15. Willis P, Candle AB, Fayrer-Hosken RA. Equine oocyte in vitro maturation: Influence of sera, time and hormones. Molec Reprod Devel 1991; 30:360-368. 16. Willis P, Candle AB, Fayrer-Hosken RA. Ultrastructure of equine oocytes matured in vitro. Theriogenology 1993; 39:340 abstr. 17. Zhang JJ, Boyle MS, Allen WR, Galli C. Recent studies on in vivo fertilization of in vitro matured horse oocytes. Equine Vet J 1989; 8 (Suppl): 101-104.