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Confocal microscopy of germinal vesicle-stage equine oocytes
ELSEVIER
CONFOCAL MICROSCOPY OF GERMINAL VESICLE-STAGE EQUINE OOCYTES T.L. Gable and G.L. Woodsa Northwest Equine Reproduction Laboratory, Department of Animal and Veterinary Science University of Idaho, Moscow, Idaho 83844-2201, USA Received for publication: July 8, 1999 Accepted: January 5, 2000 ABSTRACT The objectives were to compare cumulus type with nucleus form in equine cumulus oocyte complexes (COCs), to define the percentage of germinal vesicle (GV)-stage oocytes within a population of mares, and to further define GV nucleus shapes of equine oocytes. Cumulus types were as follows: 1) compact (56/208, 26.9%), 2) slightly expanded (37/208, 17.8%), 3) moderately expanded (27/208, 13.0%), 4) greatly expanded (15/208, 7.2%), or 5) denuded (73/208, 35.1%). One hundred thirty of 208 COCs (62.5%) were GV-stage, 21/208 (10.1%) were condensed chromatin-stage, 8/208 (3.8%) were polar body-stage, 40/208 (19.2%) were negative (nonstaining), and 9/208 (4.3 %) were fragmented. Cumulus types were associated with nucleus forms because higher proportions (P < 0.05) of GV-stage oocytes occurred in compact (42/56, 75.0%), slightly expanded (30/37, 81.1%), moderately expanded (16/27, 59.3%), or denuded (40/73, 54.8%) COCs than in greatly expanded (2/15, 13.3%) COCs. In contrast, lower proportions (P < 0.05) of condensed chromatin-stage oocytes occurred in compact (3/56, 5.4%), slightly expanded (0/37, 0.0%), moderately expanded (3/27, 11.1%), or denuded (9/73, 12.3%) COCs than in greatly expanded (6/15, 40.0%) COCs, and lower proportions (P < 0.05) of polar body-stage oocytes occurred in compact (0/56, 0.0%) or denuded (2/73, 2.7%) COCs than in greatly expanded (3/15, 20.0%) COCs. Germinal vesicle-stage equine oocytes had 4 distinct shapes, with higher proportions (P < 0.05) having large-regular (54/130, 41.5 %) than scattered (10/130, 7.7 %), small-round (29/130, 22.3 %), or large-irregular (37/130, 28.5 %) shapes. Lower proportions (P < 0.05) of large-regular GVs occurred in compact (11/42, 26.2 %) COCs than in slightly expanded (15/30, 50.0%), or moderately expanded (12/16, 75.0%) COCs. Therefore oocytes with the large-regular GV shape are probably more advanced in development. © 2001 by Elsevier Science Inc.
Key words: equine, oocyte, cumulus, germinal vesicle Acknowledgments The authors thank Don Jacklin, Marcelo Miragaya, Larry Olsen, Javier Aguilar, Elaine Crossley, and Ann Norton. This research was supported by the Idaho Equine Education Bill. This is Idaho Agricultural Experiment Station Publication No. 00A01. aCorrespondence and reprint requests: G.L. Woods, Northwest Equine Reproduction Laboratory, Department of Animal and Veterinary Science, University of Idaho, Moscow, Idaho 83844-2201; FAX: (208) 885-0501. Theriogenology 55:1417-1430, 2001 © 2001 Elsevier Science Inc.
0093-691)(/01/S-see front matter PII: S0093-691X(01)00491-5
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Theriogenology INTRODUCTION
In vitro oocyte maturation (IVM) rates are lower for mares (15 to 65% [1-5, 7, 12, 14, 15, 17, 20, 24, 27]) than for cows (85% [26]), ewes (86% [11]), sows (90% [9]), or women (70% [25]). One potential reason for the low IVM rates in mares is that oocytes of different, unknown developmental stages are subjected to the same IVM treatment. To improve IVM rates, oocytes of a similar developmental stage (e.g., GV stage) should be cultured together. This requires a clear definition of the starting population of oocytes, so that similar-stage oocytes can be selected for specific I V M protocols. First, the morphologies of equine oocytes at the time of their recovery should be defined, beginning with the cumulus. If the degree of cumulus expansion of equine COCs is associated with GV nucleus stage, COCs with less cumulus expansion would be expected to yield a higher percentage of GV-stage oocytes. However, Aim and Hinrichs (1) and Hinrichs et al. (14, 15) detected no greater percentage of GV-stage oocytes from compact than from expanded equine COCs. In contrast, Choi et al, (5) and Torner and Aim (22) recovered a higher percentage of GV-stage oocytes from compact than from expanded equine COCs, and Zhang et al. (27) recovered a higher percentage of GV- and GV breakdown-stage oocytes from compact than from expanded equine COCs. Second, the percentage of GV-stage oocytes within a population of mares should be defined. Although as few as 5% and as many as 92% of recovered equine oocytes are at the GV stage, most authors report 23% to 75%, with an overall mean of 55.5 + 25.9% (1, 2, 4, 5, 7, 14-17, 20, 22, 24, 27). Third, the morphology of GV-stage oocytes should also be defined. Hinrichs, et al. (16) defined 7 morphologies of GV-stage equine oocytes after using Hoechst 33258 stain and fluorescence microscopy. The objectives of this study were as follows: 1) compare cumulus type with nucleus stage in equine COCs, 2) define the percentage of GV-stage oocytes within a population of mares, and 3) further define GV shapes of equine oocytes using propidium iodide stain and confocal microscopy. MATERIALS AND METHODS Overview Ovaries from a population of feedlot mares were collected at a slaughterhouse in July of 1998. Approximately 80% of mares were Quarter Horse-type between 14 and 17 months of age. Approximately 20% of mares were draft and mixed breeds of unknown ages and reproductive histories. Cumulus oocyte complexes were aspirated from ovarian follicles, sorted by cumulus types, denuded, fixed, and transported to the laboratory in fixative solution. Oocytes were stained with propidium iodide and scanned using confocal laser microscopy.
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Oocyte Collection, Classification, and Transport Within 1 h after slaughter, COCs were aspirated from ovarian follicles greater than 10 mm in diameter using a 16-gauge needle connected via a tubing apparatus to a vacuum pump. Vacuum pressure was 60 mL of fluid per min. During aspiration, the needle was used to scrape the internal follicular wall to dislodge COCs. Collected COCs were examined with a stereomicroscope for classification into the following cumulus types: 1) compact, 2) slightly expanded, 3) moderately expanded, 4) greatly expanded, or 5) denuded (Figure 1). After recovery and classification, COCs were placed by cumulus types into 35 x 10 mm petri dishes containing mare pre-ovulatory follicular fluid, where they remained on a warm stage at 38 ° C for up to 4 h. Follicular fluid was aspirated from pre-ovulatory follicles (greater than 30 mm in diameter), pooled, filtered (twice with vacuum filtration and 1.6-~tm filter paper, and once with a 0.22-~tm syringe filter), and frozen at -70 ° C. One to 2 h before use, follicular fluid was warmed to 38 ° C. Cumulus oocyte complexes were removed from follicular fluid and denuded with repeated pipetting in a 0.25 % trypsina in Ca-free phosphate-buffered salinea (PBS) solution. Oocytes were fixed in 0.1 M PBS a with 5.0% paraformaldehydea and 0.5% glutaraldehydea at 4 ° C for at least 24 h. Evaluation Each oocyte was individually mounted on a poly L lysine-coated cover glass, and covered with 10 ~tL of propidium iodide (5 ktg/mL propidium iodidea in PBS) for 15 min at room temperature. Propidium iodide solution was replaced with Slowfade ®, a light anti-fade solutionb, and the cover glass was sealed with an imaging chamber. Each oocyte was viewed with a Bio-Rad MRC1000 confocal laser scanning microscopec equipped with a krypton/argon mixed gas, multi-line mode laser adjusted to 568 nm wavelength. Nucleus forms of oocytes were classified as follows: 1) GV (distinct nucleus and brightly fluorescing chromatin), 2) condensed chromatin (no distinct nucleus with dense, fluorescent chromatin), 3) polar body (extruded polar body or two independent areas of bright fluorescence), 4) negative (absence of fluorescence), or 5) fragmented (disjointed ooplasm; Figure 2).
a Sigma, St. Louis, MO, USA. b Molecular Probes, Eugene, OR, USA. c Bio-Rad, Hercules, CA, USA.
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Compact
Slightly expanded
Moderately expanded
Greatly expanded
Denuded Figure 1. Cumulus morphologies of fleshly recovered equine cumulus oocyte complexes (x 60): compact (compacted corona and outer cumulus), slightly expanded (compacted corona and slightly expanded outer cumulus), moderately expanded (compacted corona and moderately expanded outer cumulus), greatly expanded (compacted corona and completely expanded outer cumulus), and denuded (with or without a corona and completely lacking an outer cumulus).
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Figure 2. Nuclear morphologies of freshly recovered equine cumulus oocyte complexes (x 220): germinal vesicle (GV; distinct nucleus and brightly fluorescing chromatin), condensed chromatin (no distinct nucleus with dense, fluorescent chromatin), polar body (extruded polar body or two independent areas of bright fluorescence), negative (absence of fluorescence), and fragmented (disjointed ooplasm).
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Statistical Analysis Proportions of compact, slightly expanded, moderately expanded, greatly expanded, and denuded COCs with germinal vesicle, condensed chromatin, polar body, negative, and fragmented nucleus forms were compared by Chi-square analyses. Additionally, proportions of compact, slightly expanded, moderately expanded, greatly expanded, and denuded COCs with each of the germinal vesicle shapes were compared by Chi-square analyses. Fisher's exact test was used if less than 5 COCs were in any one comparison group. In all cases, proportions were considered significantly different when P < 0.05. RESULTS Two hundred ninety-three COCs were recovered from 610 follicles of 114 ovaries from 57 mares. Two hundred eight of 293 COCs (71.0%) are included in this study because 63 (21.5%) were lost during processing and 22 (7.5%) could not be categorized by nucleus form. Fifty-six of 208 (26.9%) COCs were compact, 37/208 (17.8%) were slightly expanded, 27/208 (13.0%) were moderately expanded, 15/208 (7.2%) were greatly expanded, and 73/208 (35.1%) were denuded (Figure 3). One hundred thirty of 208 COCs (62.5%) were GV-stage, 21/208 (10.1%) were condensed chromatin-stage, 8/208 (3.8 %) were polar body-stage, 40/208 (19.2%) were negative, and 9/208 (4.3%) were fragmented. Higher proportions (P < 0.05) of GV-stage oocytes occurred in compact (42/56, 75.0%), slightly expanded (30/37, 81.1%), moderately expanded (16/27, 59.3%), or denuded (40/73, 54.8%) COCs than in greatly expanded (2/15, 13.3%) COCs (Figure 3). In contrast, lower proportions (P < 0.05) of condensed chromatin-stage oocytes occurred in compact (3/56, 5.4 %), slightly expanded (0/37, 0.0 %), moderately expanded (3/27, 11.1%), or denuded (9/73, 12.3%) COCs than in greatly expanded (6/15, 40.0%) COCs, and lower proportions (P < 0.05) of polar body-stage oocytes occurred in compact (0/56, 0.0%) or denuded (2/73, 2.7%) COCs than in greatly expanded (3/15, 20.0%) COCs. Proportions of polar body-stage oocytes that occurred in slightly expanded (1/37, 2.7%), moderately expanded (2/27, 7.4%), or greatly expanded COCs were not different (P > 0.05). Proportions of negative oocytes were not different (P>0.05) among COCs with different cumulus types: compact (10/56, 17.9%), slightly expanded (5/37, 13.5%), moderately expanded (5/27, 18.5%), greatly expanded (3/15, 20.0%), and denuded (17/73, 23.3 %). Also, proportions of fragmented oocytes were not different (P > 0.05) among COCs with different cumulus types: compact (1/56, 1.8%), slightly expanded (1/37, 2.7%), moderately expanded (1/27, 3.7%), greatly expanded (1/15, 6.7%), and denuded (5/73, 6.8%). Germinal vesicle-stage oocytes were classified by nucleus shapes as follows: 1) scattered, 2) small-round, 3) large-irregular, and 4) large-regular (Figure 4). More (P < 0.05) GV-stage oocytes had large-regular (54/130, 41.5 %) than scattered ( I O/130, 7.7 %), small-round (29/130, 22.3 %), or large-irregular (37/130, 28.5 %) shapes. Lower proportions (P < 0.05) of
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large-regular GVs occurred in compact (11/42, 26.2%) COCs than in slightly expanded (15/30, 50.0%) or moderately expanded (12/16, 75.0%) COCs (Figure 5). Proportions of large-regular GVs that occurred in greatly expanded COCs (2/2, 100.0%) were not different (P > 0.05) from proportions in compact, slightly expanded, moderately expanded, or denuded COCs (14/40, 35.0%). These 4 classes of GV shape were further subcategorized into 9 GV shapes (Figure 4). Oocytes with the scattered GV shape were similar in appearance. Oocytes with small-round GV shape were further subcategorized as: solid (2/29, 6.9%), hollow (6/29, 20.7%), solid chromatin with a surrounding halo (solid-halo; 11/29, 37.9%), and hollow chromatin with a surrounding halo (hollow-halo; 10/29, 34.5 %). Oocytes with large-irregular GV shape were further subcategorized as: single, large, irregularly-shaped fluorescence (single-large; 25/37, 67.6%); and single, large, irregularly-shaped fluorescence with multiple small spots (singlelarge plus multiple-small; 12/37, 32.4%). Oocytes with the large-regular GV shape were further subcategorized as: solid (39/54, 72.2%), and hollow (15/54, 27.8%). DISCUSSION Cumulus type was associated with nucleus form in freshly recovered equine COCs. Compact, slightly expanded, moderately expanded, and denuded COCs had high proportions with GV nucleus form, while greatly expanded COCs had high proportions with condensed chromatin and polar body nucleus forms. The dividing of cumulus expansion types into slightly, moderately, and greatly expanded was key to identifying a relationship between cumulus type and nucleus form. For example, when the slightly, moderately, and greatly expanded COCs of our study were combined, no significantly higher proportions (P > 0.05) of GV nucleus form were detected between compact and expanded COCs (42/56 [75.0%] versus 48/79 [60.8%], respectively). Therefore, disagreements of previous reports concerning association between cumulus type and nucleus form of equine COCs (associated [5, 22, 27] versus not associated [1, 14, 15]) may, in part, be due to different classification systems for cumulus types. Compact, slightly expanded, and moderately expanded COCs had similar incidences of GV nucleus forms (75.0%, 81.1%, and 59.3%, respectively). Therefore, these COCs may have similar culturing requirements and could be grouped together for oocyte-culturing experiments. Greatly expanded COCs had fewer GV and more condensed chromatin (40.0%) and polar body (20.0%) nucleus forms than compact COCs. Therefore, COCs with greatly expanded cumulus may have unique, shorter culture duration requirements and should be classified separately for oocyte-culturing experiments. This notion is consistent with Hinrichs et al. (15), who achieved maximum equine oocyte IVM rates for expanded COCs at 24 h and for compact COCs at 32 h; and Zhang et al. (27) who elicited maximum equine oocyte IVM rates for expanded COCs at 24 h and for compact COCs at 30 h. Additionally, when COCs were transferred to pre-ovulatory follicles and allowed to mature in vivo, ovulate, and become fertilized, more embryos were recovered from a mare that received expanded COCs than from mares that received compact or denuded COCs (13).
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Approximately one third (35.1%, 73/208) of the COCs were denuded. In relation to the overall COC population, denuded COCs had similar (P > 0.05) proportions of GV (54.8% versus 62.5 % for the overall COC population), condensed chromatin (12.3 % versus 10.1%), polar body (2.7% versus 3.8%), negative (23.3% versus 19.2%), and fragmented (6.8% versus 4.3%) forms. Therefore, we conclude that denuded COCs represent a mixture of the population of cumulus expansion types. Perhaps the denuded type of COCs is more a reflection of how COCs were manipulated than of their nucleus form. In this experiment, we used a 16-gauge needle and a vacuum pressure of 60 mL of fluid per min. Perhaps an even smaller gauge needle and/or a lower vacuum pressure would have decreased the percentage of denuded COCs. Because 54.8% of the denuded COCs were GV-stage, this substantial source of GV-stage oocytes should not be excluded from COC culturing experiments. Also, because denuded COCs have fewer cumulus cells, which play a role in oocyte development, denuded COCs should be either cultured separately from cumulus-enclosed oocytes, or cultured with cumulus-enclosed oocytes but evaluated separately from cumulus-enclosed counterparts. Approximately one fifth (19.2%, 40/208) of the COCs were negative for DNA stain, and proportions of negative oocytes were not different among different cumulus-type groups. Therefore, similar to the denuded COCs, the negative oocytes appear not to represent a distinct phase of nucleus maturation. Four percent (9/208) of the COCs were fragmented, and proportions of fragmented oocytes were not different among different cumulus-type groups. Further work with greater numbers of oocytes is needed to determine the nucleus maturity and viability of negative and fragmented oocytes. The 62.5% (130/208) incidence of GV nucleus form in all COCs appears to be slightly higher than the 55.5% average for previous studies (1, 2, 4, 5, 7, 14-17, 20, 22, 24, 27). The wide variability (55.5 +_25.9%) among reports may be due to differences in mares (i.e., ages, breeds), seasons, or methods of evaluating nucleus form. For example, nucleus forms of equine oocytes was previously evaluated by aceto-orcein stain and phase-contrast microscopy (5, 7, 20, 22), lacmoid stain and phase contrast microscopy (27), and Hoechst 33258 or Hoechst 33342 and fluorescence microscopy (1, 12, 14-17). Because of the great variability among studies in the percentage of GV-stage equine COCs, researchers conducting culture experiments should include a time-zero control group that defines the initial proportion of GV-stage oocytes. Using propidium iodide stain and confocal microscopy, 4 shapes of GV-stage equine oocytes were described: 1) scattered, 2) small-round, 3) large-irregular, and 4) large regular. The large-regular GV shape was most abundant (54/130, 41.5%), and proportions of largeregular GV-stage COCs increased with increased cumulus expansion, suggesting that this large-regular GV shape precedes GV breakdown. The 4 GV shapes were further subcategorized into 9 distinct GV shapes: 1) scattered, 2) small-round, solid; 3) small-round, hollow; 4) small-round, solid-halo; 5) small-round, hollow-halo; 6) large-irregular, single-large; 7) large-irregular, single-large plus multiple-
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small; 8) large-regular, solid; and 9) large-regular, hollow. Some of these configurations are similar to ones described in other species, and may represent distinct phases of oocyte development. For example, the halo shape has been described in GV-stage oocytes of pigs (10) and mice (8) as a "crown," a "ring," or a "horseshoe" shape. In sows (10), this shape represents the arrested phase of development (the dictyate stage of prophase). With the attainment of meiotic competence, the GV-stage sow oocyte contains chromatin arranged into distinct, brightly-staining clumps (10), resembling the scattered shape of the present study. In other reports, however, other shapes represent the meiotically competent oocyte. The smallround, hollow and large-regular, hollow shapes of the present study, which may correspond to the nucleolar encapsulation stages described for GV-stage oocytes of mice (6) and rhesus monkeys (21), may similarly coincide with formation of perinuclear microtubule organizing centers and entry into the M phase of the meiotic cycle (19, 23). These hollow shapes may represent earlier developmental stages, however, as Debey et al. (6) determined after continued observation tharthe hollow chromatin eventually collapses into a more solid mass just before GV breakdown. Therefore, the small-regular, solid and large-regular, solid shapes may represent competence for meiotic resumption and entry into the M phase. Other evidence for this is seen in ~ e GV-stage rat oocyte, as Kopecn)? et al. (18) found that the meiotically competent oocy~ exhibited a tightly compacted nucleolus-like body, indicative of small nuclear ribonucleotide protein and maternal macromolecule storage in preparation for GV breakdown. Further investigation is needed to determine the significance of the various nucleus shapes of the GV-stage equine oocyte. Future experiments involving Hoechst 33258- or 33342-staining and fluorescence microscopy would allow monitoring throughout in vitro culture, making developmental classification of the GV-stages possible. In conclusion: 1) cumulus types and nucleus forms of equine COCs are associated; 2) 62.5 % of the oocytes from this population of mares are GV-stage; and 3) GV nucleus shapes of equine oocytes are scattered, small-round, large-irregular, and large-regular, with large-regular occurring most frequently and probably being the most advanced in development. REFERENCES 1. Aim H, Hinrichs K. Effect of cycloheximide on nuclear maturation of horse oocytes and its relation to initial cumulus morphology. J Reprod Fertil 1996;107:215-220. 2. Aim H, Torner H. In vitro maturation of horse oocytes. Theriogenology 1994;42:345349. 3. Bdzard J, Mekarska A, Goudet G, Duchamp G, Palmer E. Timing of in vivo maturation of equine preovulatory oocytes after induction of ovulation and competence for in vitro maturation of immature oocytes collected simultaneously. Proc 4 th Int Symp Equine Embryo Transf Other Adv Tech 1997. Reims, France. 4. Briick I, Gr~ndahl C, I-Iost T, Greve T. In vitro maturation of equine oocytes: Effect of follicular size, cyclic stage and season. Theriogenology 1996;46:75-84. 5. Choi YH, Hochi S, Braun J, Sato K, Oguri N. In vitro maturation of equine oocytes collected by follicle aspiration and by the slicing of ovaries. Theriogenology 1993;40:959966.
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6. Debey P, Sz6116si MS, Sz6116si D, Vautier D, Girousse A, Besombes D. Competent mouse oocytes isolated from antral follicles exhibit different chromatin organization and follow different maturation dynamics. Mol Reprod Dev 1993 ;36:59-74. 7. Del Campo MR, Donoso X, Parrish J J, Ginther OJ. Selection of follicles, preculture oocyte evaluation, and duration of culture for in vitro maturation of equine oocytes. Theriogenology 1995;43:1141-1153. 8. Donahue RF. Maturation of the mouse oocyte in vitro. I. Sequence and timing of nuclear progression. J Exp Zool 1968;169:237-250. 9. Funahashi H, Cantley T, Day BN. Different hormonal requirements of pig oocytecumulus complexes during maturation in vitro. J Reprod Fertil 1994; 101:159-165. 10. Funahashi H, Cantley TC, Day BN. Synchronization of meiosis in porcine oocytes by exposure to dibutyryl cyclic adenosine monophosphate improves developmental competence following in vitro fertilization. Biol Reprod 1997;57:49-53. 11. Galli C, Moor RM. Gonadotrophin requirements for the in vitro maturation of sheep oocytes and their subsequent embryonic development. Theriogenology 1991;35:10831093. 12. Goudet G, B6zard J, Duchamp G, G6rard N, Palmer E. Equine oocyte competence for nuclear and cytoplasmic in vitro maturation: Effect of follicle size and hormonal environment. Biol Reprod 1997;57:232-245. 13. Hinrichs K, DiGiorgio LM. Embryonic development after intra-follicular transfer of horse oocytes. J Reprod Fertil 1991;44(Suppl):369-374. 14. Hinrichs K, Martin MG, Schmidt AL, Friedman PP. Effect of follicular components on meiotic arrest and resumption in horse oocytes. J Reprod Fertil 1995; 104:149-156. 15. Hinrichs K, Schmidt AL, Friedman PP, Selgrath JP, Martin MG. In vitro maturation of horse oocytes: Characterization of chromatin configuration using fluorescence microscopy. Biol Reprod 1993;48:363-370. 16. Hinrichs K, Schmidt AL, Selgrath JP. Atlas of chromatin configurations of germinal vesicle-stage and maturing horse oocytes. Equine Vet J 1993; 15(Suppl):60-63. 17. Hinrichs K, Williams, KA. Relationships among oocyte-cumulus morphology, follicular atresia, initial chromatin configuration, and oocyte meiotic competence in the horse. Biol Reprod 1997;57:377-384. 18. Kopecn~, V, Landa V, Malatesta M, Martin TE, Fakan S. Immunoelectron microscope analyses of rat germinal vesicle-stage oocyte nucleolus-like bodies. Reprod Nutr Dev 1996;36:667-679. 19. Mattson BA, Albertini DE. Oogenesis: chromatin and microtubule dynamics during meiotic prophase [mouse]. Mol Reprod Dev 1989;25:374-383. 20. Okolski A, B~zard J, Magistrini M, Palmer E. Maturation of oocytes from normal and atretic equine ovarian follicles as affected by steroid concentrations. J Reprod Fertil 1991 ;44(Suppl): 385-392. 21. Schramm RD, Tennier MT, Boatman DE, Bavister BD. Chromatin configurations and meiotic competence of oocytes are related to follicular diameter in nonstimulated rhesus monkeys. Biol Reprod 1993;48:349-356. 22. Torner H, Aim H. Meiotic configuration of horse oocytes in relation to the morphology of the cumulus-oocyte complex. Biol Reprod Mono 1 1995:253-259.
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23. Wickramasinghe D, Ebert KM, Albertini DF. Meiotic competence acquisition is associated with the appearance of M-phase characteristics in growing mouse oocytes. Dev Biol 1991;143:162-172. 24. Willis P, Caudle AB, Fayrer-Hosken RA. Fine structure of equine oocytes matured in vitro for 15 hours. Mol Reprod Dev 1994;37:87-92. 25. Wynn P, Picton HM, Krapez JA, Rutherford AJ, Balen AH, Gosden RG. Pretreatment with follicle stimulating hormone promotes the numbers of human oocytes reaching metaphase II by in-vitro maturation. Hum Reprod 1998;13:3132-3138. 26. Younis AI, Brackett BG, Fayrer-Hosken RA. Influence of serum and hormones on bovine • oocyte maturation and fertilization in vitro. Gamete Res 1989;23:189-201. 27. Zhang JJ, Boyle MS, Allen WR, Galli C. Recent studies on in vivo fertilisation if in vitro matured horse oocytes. Equine Vet J 1989;8(Suppl): 101-104.
CONFOCAL MICROSCOPY OF GERMINAL VESICLE-STAGE EQUINE OOCYTES T.L. Gable and G.L. Woodsa Northwest Equine Reproduction Laboratory, Department of Animal and Veterinary Science University of Idaho, Moscow, Idaho 83844-2201, USA Received for publication: July 8, 1999 Accepted: January 5, 2000 ABSTRACT The objectives were to compare cumulus type with nucleus form in equine cumulus oocyte complexes (COCs), to define the percentage of germinal vesicle (GV)-stage oocytes within a population of mares, and to further define GV nucleus shapes of equine oocytes. Cumulus types were as follows: 1) compact (56/208, 26.9%), 2) slightly expanded (37/208, 17.8%), 3) moderately expanded (27/208, 13.0%), 4) greatly expanded (15/208, 7.2%), or 5) denuded (73/208, 35.1%). One hundred thirty of 208 COCs (62.5%) were GV-stage, 21/208 (10.1%) were condensed chromatin-stage, 8/208 (3.8%) were polar body-stage, 40/208 (19.2%) were negative (nonstaining), and 9/208 (4.3 %) were fragmented. Cumulus types were associated with nucleus forms because higher proportions (P < 0.05) of GV-stage oocytes occurred in compact (42/56, 75.0%), slightly expanded (30/37, 81.1%), moderately expanded (16/27, 59.3%), or denuded (40/73, 54.8%) COCs than in greatly expanded (2/15, 13.3%) COCs. In contrast, lower proportions (P < 0.05) of condensed chromatin-stage oocytes occurred in compact (3/56, 5.4%), slightly expanded (0/37, 0.0%), moderately expanded (3/27, 11.1%), or denuded (9/73, 12.3%) COCs than in greatly expanded (6/15, 40.0%) COCs, and lower proportions (P < 0.05) of polar body-stage oocytes occurred in compact (0/56, 0.0%) or denuded (2/73, 2.7%) COCs than in greatly expanded (3/15, 20.0%) COCs. Germinal vesicle-stage equine oocytes had 4 distinct shapes, with higher proportions (P < 0.05) having large-regular (54/130, 41.5 %) than scattered (10/130, 7.7 %), small-round (29/130, 22.3 %), or large-irregular (37/130, 28.5 %) shapes. Lower proportions (P < 0.05) of large-regular GVs occurred in compact (11/42, 26.2 %) COCs than in slightly expanded (15/30, 50.0%), or moderately expanded (12/16, 75.0%) COCs. Therefore oocytes with the large-regular GV shape are probably more advanced in development. © 2001 by Elsevier Science Inc.
Key words: equine, oocyte, cumulus, germinal vesicle Acknowledgments The authors thank Don Jacklin, Marcelo Miragaya, Larry Olsen, Javier Aguilar, Elaine Crossley, and Ann Norton. This research was supported by the Idaho Equine Education Bill. This is Idaho Agricultural Experiment Station Publication No. 00A01. aCorrespondence and reprint requests: G.L. Woods, Northwest Equine Reproduction Laboratory, Department of Animal and Veterinary Science, University of Idaho, Moscow, Idaho 83844-2201; FAX: (208) 885-0501. Theriogenology 55:1417-1430, 2001 © 2001 Elsevier Science Inc.
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Theriogenology INTRODUCTION
In vitro oocyte maturation (IVM) rates are lower for mares (15 to 65% [1-5, 7, 12, 14, 15, 17, 20, 24, 27]) than for cows (85% [26]), ewes (86% [11]), sows (90% [9]), or women (70% [25]). One potential reason for the low IVM rates in mares is that oocytes of different, unknown developmental stages are subjected to the same IVM treatment. To improve IVM rates, oocytes of a similar developmental stage (e.g., GV stage) should be cultured together. This requires a clear definition of the starting population of oocytes, so that similar-stage oocytes can be selected for specific I V M protocols. First, the morphologies of equine oocytes at the time of their recovery should be defined, beginning with the cumulus. If the degree of cumulus expansion of equine COCs is associated with GV nucleus stage, COCs with less cumulus expansion would be expected to yield a higher percentage of GV-stage oocytes. However, Aim and Hinrichs (1) and Hinrichs et al. (14, 15) detected no greater percentage of GV-stage oocytes from compact than from expanded equine COCs. In contrast, Choi et al, (5) and Torner and Aim (22) recovered a higher percentage of GV-stage oocytes from compact than from expanded equine COCs, and Zhang et al. (27) recovered a higher percentage of GV- and GV breakdown-stage oocytes from compact than from expanded equine COCs. Second, the percentage of GV-stage oocytes within a population of mares should be defined. Although as few as 5% and as many as 92% of recovered equine oocytes are at the GV stage, most authors report 23% to 75%, with an overall mean of 55.5 + 25.9% (1, 2, 4, 5, 7, 14-17, 20, 22, 24, 27). Third, the morphology of GV-stage oocytes should also be defined. Hinrichs, et al. (16) defined 7 morphologies of GV-stage equine oocytes after using Hoechst 33258 stain and fluorescence microscopy. The objectives of this study were as follows: 1) compare cumulus type with nucleus stage in equine COCs, 2) define the percentage of GV-stage oocytes within a population of mares, and 3) further define GV shapes of equine oocytes using propidium iodide stain and confocal microscopy. MATERIALS AND METHODS Overview Ovaries from a population of feedlot mares were collected at a slaughterhouse in July of 1998. Approximately 80% of mares were Quarter Horse-type between 14 and 17 months of age. Approximately 20% of mares were draft and mixed breeds of unknown ages and reproductive histories. Cumulus oocyte complexes were aspirated from ovarian follicles, sorted by cumulus types, denuded, fixed, and transported to the laboratory in fixative solution. Oocytes were stained with propidium iodide and scanned using confocal laser microscopy.
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Oocyte Collection, Classification, and Transport Within 1 h after slaughter, COCs were aspirated from ovarian follicles greater than 10 mm in diameter using a 16-gauge needle connected via a tubing apparatus to a vacuum pump. Vacuum pressure was 60 mL of fluid per min. During aspiration, the needle was used to scrape the internal follicular wall to dislodge COCs. Collected COCs were examined with a stereomicroscope for classification into the following cumulus types: 1) compact, 2) slightly expanded, 3) moderately expanded, 4) greatly expanded, or 5) denuded (Figure 1). After recovery and classification, COCs were placed by cumulus types into 35 x 10 mm petri dishes containing mare pre-ovulatory follicular fluid, where they remained on a warm stage at 38 ° C for up to 4 h. Follicular fluid was aspirated from pre-ovulatory follicles (greater than 30 mm in diameter), pooled, filtered (twice with vacuum filtration and 1.6-~tm filter paper, and once with a 0.22-~tm syringe filter), and frozen at -70 ° C. One to 2 h before use, follicular fluid was warmed to 38 ° C. Cumulus oocyte complexes were removed from follicular fluid and denuded with repeated pipetting in a 0.25 % trypsina in Ca-free phosphate-buffered salinea (PBS) solution. Oocytes were fixed in 0.1 M PBS a with 5.0% paraformaldehydea and 0.5% glutaraldehydea at 4 ° C for at least 24 h. Evaluation Each oocyte was individually mounted on a poly L lysine-coated cover glass, and covered with 10 ~tL of propidium iodide (5 ktg/mL propidium iodidea in PBS) for 15 min at room temperature. Propidium iodide solution was replaced with Slowfade ®, a light anti-fade solutionb, and the cover glass was sealed with an imaging chamber. Each oocyte was viewed with a Bio-Rad MRC1000 confocal laser scanning microscopec equipped with a krypton/argon mixed gas, multi-line mode laser adjusted to 568 nm wavelength. Nucleus forms of oocytes were classified as follows: 1) GV (distinct nucleus and brightly fluorescing chromatin), 2) condensed chromatin (no distinct nucleus with dense, fluorescent chromatin), 3) polar body (extruded polar body or two independent areas of bright fluorescence), 4) negative (absence of fluorescence), or 5) fragmented (disjointed ooplasm; Figure 2).
a Sigma, St. Louis, MO, USA. b Molecular Probes, Eugene, OR, USA. c Bio-Rad, Hercules, CA, USA.
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Compact
Slightly expanded
Moderately expanded
Greatly expanded
Denuded Figure 1. Cumulus morphologies of fleshly recovered equine cumulus oocyte complexes (x 60): compact (compacted corona and outer cumulus), slightly expanded (compacted corona and slightly expanded outer cumulus), moderately expanded (compacted corona and moderately expanded outer cumulus), greatly expanded (compacted corona and completely expanded outer cumulus), and denuded (with or without a corona and completely lacking an outer cumulus).
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GV
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Figure 2. Nuclear morphologies of freshly recovered equine cumulus oocyte complexes (x 220): germinal vesicle (GV; distinct nucleus and brightly fluorescing chromatin), condensed chromatin (no distinct nucleus with dense, fluorescent chromatin), polar body (extruded polar body or two independent areas of bright fluorescence), negative (absence of fluorescence), and fragmented (disjointed ooplasm).
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Statistical Analysis Proportions of compact, slightly expanded, moderately expanded, greatly expanded, and denuded COCs with germinal vesicle, condensed chromatin, polar body, negative, and fragmented nucleus forms were compared by Chi-square analyses. Additionally, proportions of compact, slightly expanded, moderately expanded, greatly expanded, and denuded COCs with each of the germinal vesicle shapes were compared by Chi-square analyses. Fisher's exact test was used if less than 5 COCs were in any one comparison group. In all cases, proportions were considered significantly different when P < 0.05. RESULTS Two hundred ninety-three COCs were recovered from 610 follicles of 114 ovaries from 57 mares. Two hundred eight of 293 COCs (71.0%) are included in this study because 63 (21.5%) were lost during processing and 22 (7.5%) could not be categorized by nucleus form. Fifty-six of 208 (26.9%) COCs were compact, 37/208 (17.8%) were slightly expanded, 27/208 (13.0%) were moderately expanded, 15/208 (7.2%) were greatly expanded, and 73/208 (35.1%) were denuded (Figure 3). One hundred thirty of 208 COCs (62.5%) were GV-stage, 21/208 (10.1%) were condensed chromatin-stage, 8/208 (3.8 %) were polar body-stage, 40/208 (19.2%) were negative, and 9/208 (4.3%) were fragmented. Higher proportions (P < 0.05) of GV-stage oocytes occurred in compact (42/56, 75.0%), slightly expanded (30/37, 81.1%), moderately expanded (16/27, 59.3%), or denuded (40/73, 54.8%) COCs than in greatly expanded (2/15, 13.3%) COCs (Figure 3). In contrast, lower proportions (P < 0.05) of condensed chromatin-stage oocytes occurred in compact (3/56, 5.4 %), slightly expanded (0/37, 0.0 %), moderately expanded (3/27, 11.1%), or denuded (9/73, 12.3%) COCs than in greatly expanded (6/15, 40.0%) COCs, and lower proportions (P < 0.05) of polar body-stage oocytes occurred in compact (0/56, 0.0%) or denuded (2/73, 2.7%) COCs than in greatly expanded (3/15, 20.0%) COCs. Proportions of polar body-stage oocytes that occurred in slightly expanded (1/37, 2.7%), moderately expanded (2/27, 7.4%), or greatly expanded COCs were not different (P > 0.05). Proportions of negative oocytes were not different (P>0.05) among COCs with different cumulus types: compact (10/56, 17.9%), slightly expanded (5/37, 13.5%), moderately expanded (5/27, 18.5%), greatly expanded (3/15, 20.0%), and denuded (17/73, 23.3 %). Also, proportions of fragmented oocytes were not different (P > 0.05) among COCs with different cumulus types: compact (1/56, 1.8%), slightly expanded (1/37, 2.7%), moderately expanded (1/27, 3.7%), greatly expanded (1/15, 6.7%), and denuded (5/73, 6.8%). Germinal vesicle-stage oocytes were classified by nucleus shapes as follows: 1) scattered, 2) small-round, 3) large-irregular, and 4) large-regular (Figure 4). More (P < 0.05) GV-stage oocytes had large-regular (54/130, 41.5 %) than scattered ( I O/130, 7.7 %), small-round (29/130, 22.3 %), or large-irregular (37/130, 28.5 %) shapes. Lower proportions (P < 0.05) of
Theriogenology
1423
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large-regular GVs occurred in compact (11/42, 26.2%) COCs than in slightly expanded (15/30, 50.0%) or moderately expanded (12/16, 75.0%) COCs (Figure 5). Proportions of large-regular GVs that occurred in greatly expanded COCs (2/2, 100.0%) were not different (P > 0.05) from proportions in compact, slightly expanded, moderately expanded, or denuded COCs (14/40, 35.0%). These 4 classes of GV shape were further subcategorized into 9 GV shapes (Figure 4). Oocytes with the scattered GV shape were similar in appearance. Oocytes with small-round GV shape were further subcategorized as: solid (2/29, 6.9%), hollow (6/29, 20.7%), solid chromatin with a surrounding halo (solid-halo; 11/29, 37.9%), and hollow chromatin with a surrounding halo (hollow-halo; 10/29, 34.5 %). Oocytes with large-irregular GV shape were further subcategorized as: single, large, irregularly-shaped fluorescence (single-large; 25/37, 67.6%); and single, large, irregularly-shaped fluorescence with multiple small spots (singlelarge plus multiple-small; 12/37, 32.4%). Oocytes with the large-regular GV shape were further subcategorized as: solid (39/54, 72.2%), and hollow (15/54, 27.8%). DISCUSSION Cumulus type was associated with nucleus form in freshly recovered equine COCs. Compact, slightly expanded, moderately expanded, and denuded COCs had high proportions with GV nucleus form, while greatly expanded COCs had high proportions with condensed chromatin and polar body nucleus forms. The dividing of cumulus expansion types into slightly, moderately, and greatly expanded was key to identifying a relationship between cumulus type and nucleus form. For example, when the slightly, moderately, and greatly expanded COCs of our study were combined, no significantly higher proportions (P > 0.05) of GV nucleus form were detected between compact and expanded COCs (42/56 [75.0%] versus 48/79 [60.8%], respectively). Therefore, disagreements of previous reports concerning association between cumulus type and nucleus form of equine COCs (associated [5, 22, 27] versus not associated [1, 14, 15]) may, in part, be due to different classification systems for cumulus types. Compact, slightly expanded, and moderately expanded COCs had similar incidences of GV nucleus forms (75.0%, 81.1%, and 59.3%, respectively). Therefore, these COCs may have similar culturing requirements and could be grouped together for oocyte-culturing experiments. Greatly expanded COCs had fewer GV and more condensed chromatin (40.0%) and polar body (20.0%) nucleus forms than compact COCs. Therefore, COCs with greatly expanded cumulus may have unique, shorter culture duration requirements and should be classified separately for oocyte-culturing experiments. This notion is consistent with Hinrichs et al. (15), who achieved maximum equine oocyte IVM rates for expanded COCs at 24 h and for compact COCs at 32 h; and Zhang et al. (27) who elicited maximum equine oocyte IVM rates for expanded COCs at 24 h and for compact COCs at 30 h. Additionally, when COCs were transferred to pre-ovulatory follicles and allowed to mature in vivo, ovulate, and become fertilized, more embryos were recovered from a mare that received expanded COCs than from mares that received compact or denuded COCs (13).
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1426
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Approximately one third (35.1%, 73/208) of the COCs were denuded. In relation to the overall COC population, denuded COCs had similar (P > 0.05) proportions of GV (54.8% versus 62.5 % for the overall COC population), condensed chromatin (12.3 % versus 10.1%), polar body (2.7% versus 3.8%), negative (23.3% versus 19.2%), and fragmented (6.8% versus 4.3%) forms. Therefore, we conclude that denuded COCs represent a mixture of the population of cumulus expansion types. Perhaps the denuded type of COCs is more a reflection of how COCs were manipulated than of their nucleus form. In this experiment, we used a 16-gauge needle and a vacuum pressure of 60 mL of fluid per min. Perhaps an even smaller gauge needle and/or a lower vacuum pressure would have decreased the percentage of denuded COCs. Because 54.8% of the denuded COCs were GV-stage, this substantial source of GV-stage oocytes should not be excluded from COC culturing experiments. Also, because denuded COCs have fewer cumulus cells, which play a role in oocyte development, denuded COCs should be either cultured separately from cumulus-enclosed oocytes, or cultured with cumulus-enclosed oocytes but evaluated separately from cumulus-enclosed counterparts. Approximately one fifth (19.2%, 40/208) of the COCs were negative for DNA stain, and proportions of negative oocytes were not different among different cumulus-type groups. Therefore, similar to the denuded COCs, the negative oocytes appear not to represent a distinct phase of nucleus maturation. Four percent (9/208) of the COCs were fragmented, and proportions of fragmented oocytes were not different among different cumulus-type groups. Further work with greater numbers of oocytes is needed to determine the nucleus maturity and viability of negative and fragmented oocytes. The 62.5% (130/208) incidence of GV nucleus form in all COCs appears to be slightly higher than the 55.5% average for previous studies (1, 2, 4, 5, 7, 14-17, 20, 22, 24, 27). The wide variability (55.5 +_25.9%) among reports may be due to differences in mares (i.e., ages, breeds), seasons, or methods of evaluating nucleus form. For example, nucleus forms of equine oocytes was previously evaluated by aceto-orcein stain and phase-contrast microscopy (5, 7, 20, 22), lacmoid stain and phase contrast microscopy (27), and Hoechst 33258 or Hoechst 33342 and fluorescence microscopy (1, 12, 14-17). Because of the great variability among studies in the percentage of GV-stage equine COCs, researchers conducting culture experiments should include a time-zero control group that defines the initial proportion of GV-stage oocytes. Using propidium iodide stain and confocal microscopy, 4 shapes of GV-stage equine oocytes were described: 1) scattered, 2) small-round, 3) large-irregular, and 4) large regular. The large-regular GV shape was most abundant (54/130, 41.5%), and proportions of largeregular GV-stage COCs increased with increased cumulus expansion, suggesting that this large-regular GV shape precedes GV breakdown. The 4 GV shapes were further subcategorized into 9 distinct GV shapes: 1) scattered, 2) small-round, solid; 3) small-round, hollow; 4) small-round, solid-halo; 5) small-round, hollow-halo; 6) large-irregular, single-large; 7) large-irregular, single-large plus multiple-
Theriogenology
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small; 8) large-regular, solid; and 9) large-regular, hollow. Some of these configurations are similar to ones described in other species, and may represent distinct phases of oocyte development. For example, the halo shape has been described in GV-stage oocytes of pigs (10) and mice (8) as a "crown," a "ring," or a "horseshoe" shape. In sows (10), this shape represents the arrested phase of development (the dictyate stage of prophase). With the attainment of meiotic competence, the GV-stage sow oocyte contains chromatin arranged into distinct, brightly-staining clumps (10), resembling the scattered shape of the present study. In other reports, however, other shapes represent the meiotically competent oocyte. The smallround, hollow and large-regular, hollow shapes of the present study, which may correspond to the nucleolar encapsulation stages described for GV-stage oocytes of mice (6) and rhesus monkeys (21), may similarly coincide with formation of perinuclear microtubule organizing centers and entry into the M phase of the meiotic cycle (19, 23). These hollow shapes may represent earlier developmental stages, however, as Debey et al. (6) determined after continued observation tharthe hollow chromatin eventually collapses into a more solid mass just before GV breakdown. Therefore, the small-regular, solid and large-regular, solid shapes may represent competence for meiotic resumption and entry into the M phase. Other evidence for this is seen in ~ e GV-stage rat oocyte, as Kopecn)? et al. (18) found that the meiotically competent oocy~ exhibited a tightly compacted nucleolus-like body, indicative of small nuclear ribonucleotide protein and maternal macromolecule storage in preparation for GV breakdown. Further investigation is needed to determine the significance of the various nucleus shapes of the GV-stage equine oocyte. Future experiments involving Hoechst 33258- or 33342-staining and fluorescence microscopy would allow monitoring throughout in vitro culture, making developmental classification of the GV-stages possible. In conclusion: 1) cumulus types and nucleus forms of equine COCs are associated; 2) 62.5 % of the oocytes from this population of mares are GV-stage; and 3) GV nucleus shapes of equine oocytes are scattered, small-round, large-irregular, and large-regular, with large-regular occurring most frequently and probably being the most advanced in development. REFERENCES 1. Aim H, Hinrichs K. Effect of cycloheximide on nuclear maturation of horse oocytes and its relation to initial cumulus morphology. J Reprod Fertil 1996;107:215-220. 2. Aim H, Torner H. In vitro maturation of horse oocytes. Theriogenology 1994;42:345349. 3. Bdzard J, Mekarska A, Goudet G, Duchamp G, Palmer E. Timing of in vivo maturation of equine preovulatory oocytes after induction of ovulation and competence for in vitro maturation of immature oocytes collected simultaneously. Proc 4 th Int Symp Equine Embryo Transf Other Adv Tech 1997. Reims, France. 4. Briick I, Gr~ndahl C, I-Iost T, Greve T. In vitro maturation of equine oocytes: Effect of follicular size, cyclic stage and season. Theriogenology 1996;46:75-84. 5. Choi YH, Hochi S, Braun J, Sato K, Oguri N. In vitro maturation of equine oocytes collected by follicle aspiration and by the slicing of ovaries. Theriogenology 1993;40:959966.
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6. Debey P, Sz6116si MS, Sz6116si D, Vautier D, Girousse A, Besombes D. Competent mouse oocytes isolated from antral follicles exhibit different chromatin organization and follow different maturation dynamics. Mol Reprod Dev 1993 ;36:59-74. 7. Del Campo MR, Donoso X, Parrish J J, Ginther OJ. Selection of follicles, preculture oocyte evaluation, and duration of culture for in vitro maturation of equine oocytes. Theriogenology 1995;43:1141-1153. 8. Donahue RF. Maturation of the mouse oocyte in vitro. I. Sequence and timing of nuclear progression. J Exp Zool 1968;169:237-250. 9. Funahashi H, Cantley T, Day BN. Different hormonal requirements of pig oocytecumulus complexes during maturation in vitro. J Reprod Fertil 1994; 101:159-165. 10. Funahashi H, Cantley TC, Day BN. Synchronization of meiosis in porcine oocytes by exposure to dibutyryl cyclic adenosine monophosphate improves developmental competence following in vitro fertilization. Biol Reprod 1997;57:49-53. 11. Galli C, Moor RM. Gonadotrophin requirements for the in vitro maturation of sheep oocytes and their subsequent embryonic development. Theriogenology 1991;35:10831093. 12. Goudet G, B6zard J, Duchamp G, G6rard N, Palmer E. Equine oocyte competence for nuclear and cytoplasmic in vitro maturation: Effect of follicle size and hormonal environment. Biol Reprod 1997;57:232-245. 13. Hinrichs K, DiGiorgio LM. Embryonic development after intra-follicular transfer of horse oocytes. J Reprod Fertil 1991;44(Suppl):369-374. 14. Hinrichs K, Martin MG, Schmidt AL, Friedman PP. Effect of follicular components on meiotic arrest and resumption in horse oocytes. J Reprod Fertil 1995; 104:149-156. 15. Hinrichs K, Schmidt AL, Friedman PP, Selgrath JP, Martin MG. In vitro maturation of horse oocytes: Characterization of chromatin configuration using fluorescence microscopy. Biol Reprod 1993;48:363-370. 16. Hinrichs K, Schmidt AL, Selgrath JP. Atlas of chromatin configurations of germinal vesicle-stage and maturing horse oocytes. Equine Vet J 1993; 15(Suppl):60-63. 17. Hinrichs K, Williams, KA. Relationships among oocyte-cumulus morphology, follicular atresia, initial chromatin configuration, and oocyte meiotic competence in the horse. Biol Reprod 1997;57:377-384. 18. Kopecn~, V, Landa V, Malatesta M, Martin TE, Fakan S. Immunoelectron microscope analyses of rat germinal vesicle-stage oocyte nucleolus-like bodies. Reprod Nutr Dev 1996;36:667-679. 19. Mattson BA, Albertini DE. Oogenesis: chromatin and microtubule dynamics during meiotic prophase [mouse]. Mol Reprod Dev 1989;25:374-383. 20. Okolski A, B~zard J, Magistrini M, Palmer E. Maturation of oocytes from normal and atretic equine ovarian follicles as affected by steroid concentrations. J Reprod Fertil 1991 ;44(Suppl): 385-392. 21. Schramm RD, Tennier MT, Boatman DE, Bavister BD. Chromatin configurations and meiotic competence of oocytes are related to follicular diameter in nonstimulated rhesus monkeys. Biol Reprod 1993;48:349-356. 22. Torner H, Aim H. Meiotic configuration of horse oocytes in relation to the morphology of the cumulus-oocyte complex. Biol Reprod Mono 1 1995:253-259.
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23. Wickramasinghe D, Ebert KM, Albertini DF. Meiotic competence acquisition is associated with the appearance of M-phase characteristics in growing mouse oocytes. Dev Biol 1991;143:162-172. 24. Willis P, Caudle AB, Fayrer-Hosken RA. Fine structure of equine oocytes matured in vitro for 15 hours. Mol Reprod Dev 1994;37:87-92. 25. Wynn P, Picton HM, Krapez JA, Rutherford AJ, Balen AH, Gosden RG. Pretreatment with follicle stimulating hormone promotes the numbers of human oocytes reaching metaphase II by in-vitro maturation. Hum Reprod 1998;13:3132-3138. 26. Younis AI, Brackett BG, Fayrer-Hosken RA. Influence of serum and hormones on bovine • oocyte maturation and fertilization in vitro. Gamete Res 1989;23:189-201. 27. Zhang JJ, Boyle MS, Allen WR, Galli C. Recent studies on in vivo fertilisation if in vitro matured horse oocytes. Equine Vet J 1989;8(Suppl): 101-104.