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a Department of Animal Science, University of Missouri-Columbia, Columbia, Missouri 65211
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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8-cell stage) collected 12–48 h after IVF were separately fixed, stained by orcein, and examined under phase contrast microscopy. It was found that 27% of 2-cell, 74% of 3-cell, 51% of 4-cell, and 74% of 5- to 8-cell-stage embryos were abnormal in morphology. Morphological anomalies included fragmentation (no nucleus in one or more than one blastomere) and/or binucleation (two nuclei in one or more than one blastomere). In experiment 2, actin filament distribution of the embryos at 2-cell to blastocyst stages that were produced in vivo and in vitro were stained by rhodamine-phalloidin and examined by confocal microscopy. Actin filaments were distributed in all in vivo-derived embryos at the cell cortex, and at the joints of cells and perinucleus in 2- to 8-cell-stage embryos and in some cells of morulae and blastocysts. Actin filaments were also distributed in the cortex and at the joints of cells of all in vitro-produced embryos. However, only 20% of in vitro-produced embryos at 2- to 8-cell stages had perinuclear actin filaments in all blastomeres. Most in vitro-produced embryos had fewer perinuclear actin filaments or did not have perinuclear actin filaments in some blastomeres. Fragmentation and binucleate blastomeres were not observed in in vivo-derived embryos. In vivo-derived Day 5 (136.5 ± 60.4 nuclei per blastocyst) and Day 6 (164.5 ± 51.9 nuclei per blastocyst) blastocysts had significantly (p < 0.001) more cells than in vitro-produced Day 6 blastocysts (37.3 ± 11.7 nuclei per blastocyst). In experiment 3, when cytochalasin D, an inhibitor of microfilament polymerization, was included in the culture medium, it prevented 2- to 4-cell-stage embryos from developing to the blastocyst stage. These results indicate that abnormal actin filament distribution is one possible reason for abnormal embryo cleavage and small cell numbers in pig embryos produced in vitro. Culture conditions that mediate normal actin filament distribution may result in an improvement in embryo quality.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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In vitro production of pig embryos was based on the procedures reported in our previous study [2]. Briefly, oocytes were aspirated from antral follicles (3–6 mm in diameter) of ovaries collected from prepubertal gilts. After being washed 4 times with Hepes-buffered Tyrode's medium containing 0.1% (w:v) polyvinyl alcohol (Hepes-TL-PVA), each group of 50 oocytes surrounded by compact cumulus was cultured in BSA-free NCSU 23 medium supplemented with 0.57 mM cysteine (Sigma Chemical Co., St Louis, MO), 10% pig follicular fluid, 10 ng/ml epidermal growth factor, 10 IU/ml eCG (Intervet America Inc., Millsboro, DE) and 10 IU/ml hCG at 39°C and 5% CO2 in air in a 500-µl drop of the same medium. After being cultured for 22 h, oocytes were washed 3 times and then cultured in maturation medium without hormones for another 22 h.
After maturation, oocytes were separated from the enclosed cumulus by pipetting in maturation medium containing 0.02% hyaluronidase (Sigma). After being washed 3 times, cumulus-free oocytes were transferred to 50 µl of insemination medium, a modified Tris-buffered medium, consisting of 113.1 mM NaCl, 3.0 mM KCl, 7.5 mM CaCl2, 20.0 mM Tris, 11.0 mM glucose, 5.0 mM sodium-pyruvate, 2 mM caffeine, and 2 mg/ml BSA [2]. The dishes were kept in a CO2 incubator until spermatozoa were added for insemination. For IVF, 0.2-ml frozen ejaculated semen was thawed at 39°C in 10 ml of PBS containing 1 mg/ml BSA and antibiotics. After being washed 3 times, spermatozoa were resuspended with insemination medium to give a concentration of 5 x 105 cells/ml, and 50 µl of the sample was added to 50 µl of fertilization drops containing the oocytes. Six hours after insemination, oocytes were removed from fertilization drops and cultured in 500 µl of culture medium (NCSU 23 containing 4 mg/ml BSA) in a four-well culture plate until examination.
Collection of In Vivo Embryos
In vivo-derived embryos were surgically recovered from 12 naturally mated pigs on Days 2–6 (3 on Day 2, 6 on Day 4, 2 on Day 5, and 1 on Day 6) of the estrous cycle. Because experiments were focused on early embryos in the present study, more animals were used for collection of early-stage embryos. Briefly, sexually mature pigs, 6–8 mo of age and weighing 100–120 kg, were examined for signs of estrus twice daily by exposure to a sexually mature boar. On Day 0 of estrus, the animals were artificially inseminated, and oviducts or uteri were flushed with 30 ml Hepes-TL-PVA on Days 2–6 to collect embryos at various stages.
Morphologic Evaluations of Embryos Produced In Vitro (Experiment 1)
At 6 h after IVF, inseminated oocytes were pooled together, and after being washed 3 times in culture medium, the oocytes were randomly assigned to culture in 5 culture drops. Morphologic evaluations of embryos were conducted 12, 24, 30, 36, and 48 h after IVF. The embryos at different stages were separately mounted on slides and fixed in 25% (v:v) acetic alcohol, stained with 1% (w:v) orcein in 45% (v:v) acetic acid, and examined under a phase-contrast microscope. Fertilization rates were calculated by adding the cleaved embryos and one-cell oocytes that had been penetrated by spermatozoa. Among the cleaved embryos, four different morphological characteristics were observed in the present study. They were 1) morphological normality: embryos with each blastomere having one nucleus; 2) fragmentation: embryos with one or more than one blastomere having no nucleus; 3) binucleation: embryos with one or some blastomeres having two nuclei; and 4) embryos with both fragmented and binucleate blastomeres. The details of these different morphologies at 2- to 4-cell stages are shown in Figure 1A.
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Comparison of Morphology and Actin Filaments of In Vivo and In Vitro Embryos (Experiment 2)
In experiment 2, in vitro-produced embryos were collected at 36 h (2- to 5-cell stages) and 6 days (blastocyst) after IVF. In vivo embryos were collected as mentioned above. Embryos were fixed in 3.7% paraformaldehyde in PBS for 2 h at room temperature. After fixation, embryos were treated with 1% (v:v) Triton X-100 in PBS for 6 h at room temperature or overnight at 4°C and then washed twice in PBS and cultured in a blocking solution (PBS containing 2% BSA and 150 mM glycine) for 30 min at room temperature. After being washed for another hour in PBS, the embryos were stained with 10 IU/ml rhodamine-phalloidin (Molecular Probe, Eugene, OR) for 1 h at 39°C in PBS-Tween 20 (0.1%, v:v). After washing twice in PBS-Tween solution for 2 h at room temperature, the embryos were stained with 100 nM YO-Pro-1 iodide (Molecular Probe) for 5–10 min for examination of nuclear status. Finally, the embryos were mounted on slides and examined by using confocal microscopy.
Confocal microscopy was performed using a Bio-Rad MRC-600 confocal laser scanning imaging system (Richmond, CA) equipped with a krypton argon ion laser, mounted on an Optiphoto II Nikon microscope (Garden City, NY) equipped with 40x or 60x objectives. The image was obtained by repeated laser scanning (5 times during 5 sec) to improve the signal-to-noise ratio.
Effect of Cytochalasin D on Embryo Development and Microfilament Distribution (Experiment 3)
In experiment 3, in order to further examine the effect of microfilaments on embryo development, 2- to 4-cell embryos selected at 36 h after IVF were cultured in medium with or without 5 µM cytochalasin D, an inhibitor of microfilament polymerization. At 4.5 days (6 days after IVF) after culture, blastocyst formation and actin filament distribution were examined by methods described above.
Statistical Analysis
Four replicate trials were conducted for experiments 1 and 3, and all percentage data were subjected to arc sine transformation before statistical analysis. Comparisons were conducted by ANOVA. For examination of actin filament distribution, embryos at 36 h and 6 days (blastocyst) after IVF from four different experimental trials were used. Cell numbers in embryos from in vivo and in vitro procedures were directly compared by ANOVA. A value of p < 0.05 was considered to be statistically significant.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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As shown in Table 1, no statistical differences were observed in the sperm penetration rates among the examination time points from 12–48 h after IVF. At 12 h after IVF, 93% of fertilized oocytes had formed both male and female pronuclei and released the second polar body. Of fertilized oocytes, 35% were polyspermic. No cleaved embryos were observed at 12 h after IVF. Cleaved embryos were first observed 24 h (25%) after IVF, and the cleavage rate had significantly increased at 30 h (71%) after IVF and reached the highest point at 36 h (83%) after IVF. Cell stages (number of blastomeres) progressed as the culture time increased (Table 1). The detailed nucleus-blastomere relationship at 2- to 4-cell stages is shown in Figure 1A. From a total of 269 embryos examined, 109, 19, 74, and 67 were at the 2-cell, 3-cell, 4-cell, and 5- to 8-cell stages, respectively. Of the embryos examined, as shown in Figure 1B, 73% of 2-cell, 26% of 3-cell, 49% of 4-cell, and 26% of 5- to 8-cell embryos were normal in morphology; the proportion of embryos with abnormal morphology increased as number of blastomeres increased, from 27% at the 2-cell stage to 74% at the 5- to 8-cell stage. The relationship between culture duration and embryo morphology is summarized in Figure 1C. Most 2-cell- (67%) and 3-cell- (50%) stage embryos examined at 24 h were normal, but only 14% of 4-cell- and none of the 5- to 8-cell-stage embryos were normal. The proportion of normal embryos at the 4-cell stage increased as culture duration increased to 30–48 h. However, only 17% of 5- to 8-cell-stage embryos at 48 h showed normal morphology.
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Nucleus-Blastomere Relationship and Actin Filament Distribution of Embryos Produced In Vivo and In Vitro (Experiment 2)
As shown in Figure 2, there were great differences in the morphology of embryos produced in vivo and in vitro. In vivo-derived 2- and 4-cell embryos had clear blastomeres, especially at the 4-cell stage, and all blastomeres were separated. A wide perivitelline space (PVS) was observed in in vivo 2- to 4-cell embryos. After the 8-cell stage, the PVS was fully filled with blastomeres. The inner cell mass was clearly seen in in vivo blastocysts. However, the cytoplasm of in vitro-produced embryos was dark, and it was difficult to see clear blastomeres beyond the 4-cell stage. The PVS was fully filled with blastomeres even at the 2-cell stage. No obvious inner cell mass was observed in in vitro-produced blastocysts.
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Examination of the nucleus-blastomere relationship indicated that all in vivo embryos had normal morphology, with one nucleus per blastomere. However, because fragmentation and binucleate blastomeres were present in in vitro-produced embryos, the mean number of nuclei varied at each stage examined (Table 2). Moreover, the mean number of nuclei was usually less than the number of blastomeres. More nuclei were present in in vivo-derived blastocysts than in in vitro-produced blastocysts. Actin filaments were present at the cortex and at the joints of blastomeres of all in vivo- and in vitro-produced embryos (Table 2; Figs. 3 and 4). However, considerable differences existed in the perinuclear actin filament distribution. All in vivo-produced 2- to 8-cell-stage embryos and some blastomeres of morulae and blastocysts had perinuclear actin filaments (Fig. 3, E and G). However, as shown in Table 2, this type of actin filament distribution was observed only in 31% of 2-cell-, 14% of 3-cell-, and 18% of 4-cell-stage in vitro-produced embryos. Most in vitro-produced embryos had partial perinuclear actin filaments in their blastomeres (Fig. 4, B, F, and G), and some embryos (5% of 2-cell, 9% of 4-cell and 25% of 5-cell) did not have perinuclear actin filaments in their blastomeres (Fig. 4, A, C, and I). As shown in Figure 5, asynchronized cytoplasmic and nuclear division was observed in in vitro-produced 2-cell- and 3-cell-stage embryos. In these embryos, it was found that chromosomes were present in one or two blastomeres or that mitotic division had occurred but cytoplasmic division had not. These situations were not observed in in vivo-derived embryos at any stage.
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Effect of Cytochalasin D on the Blastocyst Formation (Experiment 3)
Of 139 2- to 4-cell-stage embryos cultured in medium without cytochalasin D, 81 (58.3%) developed to blastocysts, with a mean nuclear number of 41.4 ± 14.4 per blastocyst. Actin filaments were distributed at the cortex of blastocysts, as shown in Figure 3H. However, no embryos (0/135) cultured in medium with cytochalasin D developed to blastocysts. All of the embryos remained at the 2- to 4-cell stages. The nuclei of these embryos divided one or two times, thus forming binucleate (Fig. 6A) or polynucleate (Fig. 6B) embryos. No obvious actin filaments were present in the embryos cultured with cytochalasin D (Fig. 6, A' and B').
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The present study is the first detailed examination of morphology of pig embryos produced by IVM/IVF/IVD. Although 58.3% of cleaved embryos developed to the blastocyst stage under present conditions, the nuclear number in blastocysts was still low (less than one third of in vivo-derived blastocysts). We found that fragmentation was a major defect in these embryos. Fragmentation has also been frequently found in human embryos produced in vitro [14–16]. Studies in the human suggest that the best possible reason for embryonic fragmentation is an inadequate culture environment in vitro [14]. One in vitro culture condition differing from in vivo conditions is that the oxygen concentration is higher and, under such conditions, a higher concentration of reactive oxygen species (ROS) is produced in culture [17]. Yang et al. [14] found that fragmented blastomeres in human embryos had higher ROS than did normal embryos. High ROS is thought to increase membrane permeability and cause cell damage [18] and DNA fragmentation [19]. It may also inhibit or delay polymerization and depolymerization of actin filaments, inducing the cytoplasm and nucleus to divide asynchronously, which results in fragmentation or binucleation. Reduced oxygen strength has been used for IVM [20, 21], IVF [21], and IVD [22–24] in some mammals, and it seems superior to the concentration (20%) commonly used [17]. However, Machaty et al. [25] cultured in vivo pig embryos for 4 days under two levels (5% vs. 20%) of oxygen and did not find beneficial effects on blastocyst development with a 5% oxygen concentration. It is possible that in vivo embryos collected from oviducts have already developed the ability to reduce ROS while in the oviducts, thus masking the effects of oxygen level on embryo development.
In the present study, about 80% of inseminated oocytes were penetrated by sperm, and more than 90% of penetrated oocytes formed both male and female pronuclei. However, 35% of fertilized oocytes were polyspermic. From cleavage rates at 36 h and 48 h after IVF, we estimated that most fertilized oocytes that had formed both male and female pronuclei had cleaved. This was also consistent with the results in Experiment 1 that most uncleaved oocytes at 36–48 h after IVF were unfertilized. These results indicated that fertilized oocytes containing both male and female pronuclei can cleave irrespective of polyspermy or monospermy. Development of polyspermic pig oocytes has recently been demonstrated in our laboratory (unpublished data), although the rate of blastocyst formation and cell number in blastocysts were lower in polyspermic embryos than monospermic embryos. However, it is not clear that the abnormal embryos observed in this study resulted from monospermic oocytes or polyspermic oocytes. Because the in vitro-derived embryos used in this study were produced by IVM and IVF, oocyte maturation and activation in vitro may also affect developmental ability of embryos. Incomplete cytoplasmic maturation of pig oocytes matured in vitro was reported previously and included failure of the oocytes to form a male pronucleus after fertilization and inability to develop further. It was found that low male pronuclear formation was due to a low concentration of glutathione in the oocytes, and this has been overcome by supplementation of cysteine, a precursor of glutathione in culture medium [26]. Recently it was also reported that epidermal growth factor [27, 28] or coculture of pig oocytes with follicle shell pieces [3] improved cytoplasmic maturation of pig oocytes by improving male pronuclear formation [28] and subsequent embryo development [3, 27]. In the present study, we used an in vitro system that results in more than 30% of oocytes matured and inseminated to develop to blastocysts [1–3]. It seems that improved oocyte maturation conditions have improved cytoplasmic maturation and increased the rate of blastocyst formation but have not significantly increased the cell number in blastocysts. Previous studies in pigs clearly indicated that intracellular glutathione was an important factor affecting embryo development [2, 3, 27]. Glutathione has been found to regulate several important cellular functions, such as maintenance of cell and membrane integrity and redox status, regulation of protein and DNA synthesis, modulation of protein folding, and participation as a cofactor for various enzyme reactions or as a protector of the cell against oxidative stress [29]. As mentioned above, the latter may be more important during embryo development under in vitro conditions. After fertilization, intracellular glutathione concentration decreases significantly [26], and the embryos do not have the ability to synthesize glutathione until the blastocyst stage; thus the embryos may be exposed to conditions with a high concentration of ROS. Interestingly, recently it has been found that addition of glutathione during the sperm-oocyte reaction [30–32] and embryo culture [33, 34] significantly improved blastocyst formation without increasing intracellular glutathione [34]. It is possible that glutathione can also reduce ROS in the culture medium and thus protect the embryos against oxidative stress. It was also found that glutathione was present in oviduct secretions [34]. Put together, these results indicate that both intracellular and extracellular glutathione are important for normal embryo development, not only during oocyte maturation, but also during sperm-oocyte interaction and early embryo development.
Actin is a major component of the cytoskeleton. During oocyte maturation, fertilization, and embryo development, the polymerization and depolymerization of actin filaments is an important process. Early studies indicated that actin filaments were responsible for maintenance of the meiotic spindle, spindle rotation, polar body release, pronuclear migration, and mitotic cleavage [7, 8, 12, 13]. Abnormal actin filament distribution has been observed in in vitro-matured and -fertilized pig oocytes [12, 13]. In the present study, through a comparison of in vivo-derived and in vitro-produced embryos, we also found that microfilament distribution was significantly different between these two sources of embryos, especially in intracellular actin filaments. Recently it was reported that intracellular actin filament distribution was directly related to the 2-cell block in hamster [10] and rat [11] embryos. How actin filaments modulate embryo development is poorly understood. It has been reported that the actin cytoskeleton is required for normal mitochondria distribution [10], transfer of mitochondria to daughter cells during cell division [35], subcellular localization and morphology of Golgi complexes [36], and the distribution of mRNA [37]. It is clear that abnormal microfilament distribution results in abnormal cell function. Conditions, such as the composition of the culture medium [10,11], may affect polymerization and stabilization of actin filaments. The results in experiment 3 indicate that inhibition of actin filament polymerization prevents embryo division without affecting nuclear division, at least in 1- to 2-cell cycles. These results also suggest that the presence of binucleate cells observed in the present study is due to abnormal polymerization of actin filaments, which results in nuclear division without cytoplasmic division. In addition, asynchronized nuclear and cytoplasmic division (as shown in Figs. 4 and 5) indicates that culture conditions may delay the progression of polymerization and depolymerization of microfilaments. Most cells keep a large pool of G-actin (nonfilamentous actin) to maintain the ability to quickly reorganize the F-actin (filamentous actin) when subjected to environmental changes. It is not clear whether pig oocytes or embryos synthesize sufficient actin protein during oocyte maturation and early development. The examination of G-actin synthesis in pig oocytes and embryos may be helpful to determine why in vitro-produced embryos have less filamentous actin.
In summary, the present study indicates that poor developmental ability and decreased number of cells in pig embryos produced in vitro are due to abnormal cleavage during development. Abnormal cleavage starts at the first cell division. One of the most typical abnormal morphologies is embryo fragmentation. No fragmentation was observed in in vivo-derived embryos. The differences in microfilament distribution observed in this study between in vivo and in vitro embryos suggest that conditions for oocyte maturation and activation, and especially for embryo culture, which mediate more normal actin microfilament distribution may reduce embryo fragmentation and result in an improvement in quality of in vitro embryos.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Correspondence: Billy N. Day, 159 Animal Sciences Research Center, Department of Animal Science, University of Missouri-Columbia, Columbia, MO 65211. FAX: 573 884 7827; dayb{at}missouri.edu
3 Current address: Yong-Mahn Han, Korea Research Institution of Biosciences and Biotechnology, Teajon 305–600, Korea.
Accepted: November 29, 1998.
Received: October 8, 1998.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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