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a Institute of Zoology, Regent's Park, London, NW1 4RY, United Kingdom
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Marmoset oocytes were either subjected to two series of six electrical pulses (DC; 2 kV/cm and 70 µsec) or were incubated in 7% ethanol in PBS. Ninety-two percent (68 of 74) and 20% (8 of 40) of marmoset oocytes were activated by electrical stimulus or ethanol, respectively. Parthenogenetic (n = 3) or in vitro-fertilized (n = 2) embryos were transferred at the 4-cell stage to synchronized recipient female marmosets (n = 5). Progesterone, chorionic gonadotropin, and inhibin in the peripheral plasma of recipient animals were measured. After 33 days of gestation, recipient animals were perfused and the uteri were collected. The 2 females that had received biparental embryos and 2 of the 3 females that had received parthenogenetic embryos displayed biochemical and histological evidence of implantation.
This is the first report that a primate embryo comprising only parthenogenetic cells is capable of implantation. This highlights the need to scrutinize levels of parthenogenesis associated with human assisted reproductive technologies. Marmoset parthenogenones also provide a unique model for elucidating the roles of parental genomes in primate development.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Although murine parthenogenones can develop to postimplantation stages in vivo [7, 8], human parthenogenetic embryos only develop to the 8-cell stage when cultured in vitro [1]. This provides little information about the differential contribution of parental genomes to primate embryonic development. In humans, for ethical reasons, the development of parthenogenones to postimplantation stages cannot be studied as this would require transfer of genetically manipulated embryos. Full understanding of the role the parental genomes play in postimplantation human development requires a nonhuman primate model.
The common marmoset monkey (Callithrix jacchus) is a small New World primate that routinely produces 2–3 follicles per 28-day cycle [9, 10]. This species is particularly suited to studies of postimplantation development since the ovarian cycle can be synchronized using a prostaglandin F2
analogue [11], follicular aspiration has been developed [9, 10], embryo transfer is highly successful [12], and early postimplantation development in the marmoset has been described [13]. Here we report parthenogenetic activation of marmoset oocytes using ethanol and electrical stimulation, the development of marmoset parthenogenones in vitro, and the ability of parthenogenetic primate embryos to implant in vivo, after embryo transfer.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The experiments reported here were carried out using parous common marmoset monkeys that were housed at The Institute of Zoology, London, under conditions previously described [11]. The animals were housed and cared for according to U.K. Home Office regulations (Animals [Scientific Procedures] Act 1986) and kept either in breeding pairs or in family groups. The investigations described in this manuscript were conducted in accordance with the Guiding Principles for the Care and Use of Research Animals promulgated by the Society for the Study of Reproduction.
Synchronization of Oocyte Donors
Marmosets have a 27- to 29-day ovarian cycle with a follicular phase of 8–10 days and a luteal phase of 18–20 days [14]. Animals received 0.5 µg cloprostenol, a prostaglandin F2 analogue (Estrumate; Coopers Animal Health, Bristol, UK), by intramuscular injection at 0900 h between Days 15 and 24 after ovulation. This caused premature luteolysis and effectively reset the cycle to the beginning of the follicular phase [11]. At 1300 h, 7 days after cloprostenol administration, the animals received an i.m. injection of 75 IU of hCG (Chorulon; Centaur, Somerset, UK). Laparotomy was carried out between 1000 and 1300 h, 20–24 h post-hCG injection [10].
Marmoset Oocyte Collection
Oocytes were collected by follicular aspiration as previously described [10]. Briefly, female marmosets were anesthetized with alphadolone and alphaxalone (Saffan, ~2.5 ml/kg BW; Centaur). The ovaries and uterus were exteriorized by midline laparotomy. Follicles larger than 2 mm in diameter were aspirated with a pulled 1.5-mm-diameter glass capillary, broken off at 0.7- to 0.8-mm diameter, which was attached to a micrometer syringe. The follicular contents were drawn out and expelled into a 35-mm sterile Petri dish containing alpha modified minimum essential medium (MEM; Merck, Lutterworth Leics, UK) buffered with 25 mM Hepes, and supplemented with 0.05 mg/ml streptomycin sulfate, 0.06 mg/ml penicillin (Sigma Chemical Co., Poole, Dorset, UK), 1 IU/ml heparin (Monoparin; CP Pharmaceuticals Ltd., Wrexham, UK), and 1% heat-inactivated marmoset serum.
Evaluation of Oocytes
Oocytes were incubated briefly in 0.1% hyaluronidase (Sigma Chemical Co.) in Medium 2 [15] and gently pipetted with a flame-polished Pasteur pipette to remove cumulus cells. After cumulus cell removal it was possible to visualize the first polar body. Only oocytes that had extruded a first polar body and so were presumably in meiotic metaphase II were used in this study. Cumulus-free marmoset oocytes were incubated in MEM supplemented with 10% heat-inactivated marmoset serum in a humidified atmosphere of 5% CO2 in air, at 37°C, until activation stimulus, by electrical pulses or ethanol exposure, was applied.
Exposure of Marmoset Oocytes to Electrical Stimulation
Twenty-four hours after collection, oocytes were subjected to electrical pulses generated by an Electro Cell Manipulator 200 (BTX Inc., San Diego, CA) applied in a chamber consisting of 2 parallel, stainless steel, 0.5-mm diameter electrodes attached to a glass slide, 0.5 mm apart (part no. 450; BTX Inc.). The chamber was filled with 0.3 M mannitol in embryo culture-quality water, supplemented with 100 µM CaCl2 and 100 µM MgCl2 (Sigma Chemical Co.). The oocytes were subjected to two series of six electrical pulses (DC), 30 min apart. Each pulse was 2 kV/cm and of 70-µsec duration. Control oocytes (n = 19) were not exposed to the mannitol solution or to electrical stimulation.
Exposure of Marmoset Oocytes to Ethanol
Oocytes were incubated in 7% ethanol in PBS (Life Technologies, Paisley, Scotland) for 5 min at room temperature [16]. Some of these oocytes (n = 28) had been incubated with sperm as part of another study. All oocytes in this group were exposed to ethanol 3 days after collection (4 days post-hCG), when it was confirmed that oocytes had not been fertilized, by absence of pronuclei and lack of cleavage for 48 h postinsemination.
Culture of Marmoset Oocytes and Assessment of Parthenogenetic Activation
After activation stimulus was applied, all oocytes were washed thoroughly and cultured singly in ~50-µl drops of MEM supplemented with 10% heat-inactivated female marmoset serum overlaid with paraffin oil to prevent evaporation. Oocytes were incubated at 37°C in a humidified atmosphere of 5% CO2 in air.
Oocytes were examined, using an inverted Olympus OMT-2 microscope fitted with Nomarski optics, 6 and 18–20 h after activation stimulus, and daily thereafter. Pronuclear formation, polar body extrusion, and cell number were assessed at each observation. Parthenogenetic embryos were either left in culture till developmental arrest, as determined by lack of cleavage for at least 24 h, or transferred at the 4-cell stage to recipient females.
Transfer of Parthenogenetic Marmoset Embryos to Recipient Marmosets
Cloprostenol was administered to recipient animals on the same day as donor animals. Recipient females were housed singly from the time of cloprostenol administration until 6 days after embryo transfer. Recipients received hCG (75 IU) at 1000 h, 8 days after cloprostenol, and a 0.3-ml blood sample was taken from the femoral vein. A second blood sample was taken 48 h later. Ovulation was confirmed by plasma progesterone levels over 10 ng/ml. Parthenogenetic (n = 3) or biparental in vitro-fertilized (IVF) control (n = 2) embryos were transferred at the 4-cell stage to synchronized recipient female marmosets (n = 5) 3 days after hCG injection. Embryo transfers were carried out as previously described [12]. Briefly, at midline laparotomy, the uterus was exteriorized and a hole was made in the uterine fundus using a 19-gauge needle. The embryo was introduced into the fundus in a pulled glass Pasteur pipette attached by rubber tubing to a micrometer syringe. The contents of the pipette (approximately 3 µl) were gently expelled into the uterine lumen. Anesthesia and postoperative care were carried out as described previously [10].
In Vitro Fertilization of Marmoset BiparentalControl Embryos
In vitro fertilization was carried out as previously described [10, 17]. Briefly, sperm were collected by epididymal dissection. Both oocytes and sperm were incubated in MEM supplemented with 10 µM dibutyryl cAMP, 10 µM caffeine, 6 mg/100 ml penicillin, 5 mg/100 ml streptomycin sulphate, and 10% heat-inactivated male marmoset serum. Oocytes were incubated for 9–11 h, and sperm were incubated for at least 3 h and up to 7–8 h before insemination. The concentration of sperm was approximately 10–15 x 106 sperm/ml, and the insemination time ranged from 12–20 h. After insemination, oocytes were removed from the insemination media, washed, and placed in drops of MEM with 10% heat-inactivated female marmoset serum and incubated at 37°C in a humidified atmosphere of 5% CO2 in air. Cumulus cells were easily removed from oocytes by repeated pipetting with a flame-polished, pulled Pasteur pipette. Fertilization was confirmed by visualization of a second polar body and two or more pronuclei. Embryos were cultured in drops (approximately 50 µl) of MEM, supplemented with 6 mg/100 ml penicillin, 5 mg/100 ml streptomycin sulphate, and 10% heat-inactivated female marmoset serum. The drops were overlaid with paraffin oil (Merck). Embryos were observed daily and transferred to recipient females at the 4-cell stage, as described above.
Monitoring of Recipient Female Marmosets afterEmbryo Transfer
Approximately 0.5 ml of blood was drawn in a 1-ml syringe from the femoral vein of recipient female marmosets 2–3 times per week. The blood sample was centrifuged in the syringe casing at ~2500 rpm for 10 min. Plasma was aspirated using a glass Pasteur pipette and stored in plastic tubes at -20°C.
Three hormones—progesterone, marmoset CG, and immunoreactive (ir) inhibin—were measured. These are all diagnostic indicators of pregnancy in the peripheral plasma of recipient female marmosets. Progesterone levels were monitored by weekly ELISA developed for marmoset monkeys [18]. The day of ovulation (Day 0) was determined as the day before progesterone levels rose above 10 ng/ml [19]. CG was measured using a mouse Leydig cell bioassay [20, 21]. Ir-inhibin was measured as previously described [22], with some modifications. Briefly, levels of ir-inhibin were measured by RIA, using antiserum to the N-terminal sequence of the -subunit of human inhibin raised in sheep. In the original protocol, the tracer was monomeric inhibin -subunit. For this study, the 32-kD dimer of inhibin was used as the tracer.
Whole-Body Perfusion Fixation of RecipientFemale Marmosets
Recipients that had high (>50 ng/ml) progesterone until at least Day 26 were considered pregnant, and these animals were killed. The females were anesthetized with 1 ml of Saffan administered by i.m. injection. Once the animals were under general anesthesia, the peritoneum and thoracic cavity were opened. The ovaries were inspected for the presence of corpora lutea (CL). A 21-gauge butterfly catheter (Centaur) was inserted into the left ventricle of the heart and held in place with artery forceps, and the vena cava was cut posterior to the renal veins. The animal was perfused by delivering 200 ml heparinized (1000 IU/ml) PBS and then 200 ml 2.5% glutaraldehyde through the catheter. After perfusion, the uterus was dissected from the reproductive tract and stored in 2.5% glutaraldehyde until sectioning was carried out.
Sectioning and Staining of Marmoset Uteri
Sectioning was performed by the Department of Histopathology at the Royal Free Hospital School of Medicine (London, UK). Paraffin wax sections (3-µm thick) of perfused marmoset uteri were cut on the anterior-posterior axis, and every fifth section was mounted on a glass slide and stained with hematoxylin and eosin. Sections were inspected under brightfield microscopy, and relevant sections were photographed.
Statistical Analyses
The differences in the proportions of oocytes undergoing each type of development after ethanol or electrical exposure were analyzed by ANOVA (CSS: Statistica; Statsoft U.K., Letchworth, UK).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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A total of 122 oocytes were collected for this study, from 51 laparotomies. The average number of follicles per animal was 2.6, and the mean number of oocytes collected per animal was 2.4.
Visualization of Activation Events
Six hours after activation stimulus, some oocytes had undergone immediate cleavage (IC) to 2 cells. After 18–20 h, oocytes had either cleaved (IC), extruded a second polar body and formed one pronucleus (2PB,1PN), formed one pronucleus without extruding a second polar body (1PB,1PN), formed two pronuclei without extruding a second polar body (1PB,2PN), or failed to activate. These events occurred with the same timing after activation stimulus, regardless of whether ethanol or electrical stimulation was used.
Activation and Development of Marmoset Oocytes Exposed to Electrical Stimulation
Ninety-two percent (68 of 74) of marmoset oocytes were parthenogenetically activated after exposure to two series of six electrical pulses. The average cell number achieved in vitro was not significantly affected by the type of development (1PN,2PB; 2PN,1PB; 1PN,1PB, or IC) after activation (p < 0.05; Table 1). Nor was there a significant difference between the in vitro development of haploid (1PN,2PB or IC) and diploid (2PN,1PB or 1PN,1PB) parthenogenones in this treatment group (p < 0.05). Overall, the average cell number (± SEM) of electrically stimulated parthenogenones was 4.0 ± 0.3.
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Activation and Development of Marmoset Oocytes Exposed to Ethanol
Twenty percent (8 of 40) of oocytes exposed to ethanol were activated. Three types of development after activation were observed. Five (63%) activated oocytes underwent IC and had divided to 2 cells within 6–18 h of treatment; two oocytes (25%) were 2PB,1PN; and one oocyte was 1PB,2PN. Only the oocytes that underwent IC divided in culture. One of these embryos developed to 16 cells, one to 6 cells, one to 3 cells, and two to 2 cells in vitro. Overall, the average cell number (± SEM) of ethanol-activated parthenogenones was 4.0 ± 1.7. This was not significantly different from the average cell number reached by electrically stimulated parthenogenetic embryos.
None of the control oocytes (0 of 19) were parthenogenetically activated.
Progesterone Profiles of Recipient Marmosets
Normal serum progesterone profiles for pregnant and nonpregnant marmosets have been reported previously [23]. Briefly, in a normal nonpregnant cycle, progesterone levels rise above 10 ng/ml on the day after ovulation, increase to 50–100 ng/ml until 16–18 days after ovulation (the end of the luteal phase), and drop to baseline (<10 ng/ml) throughout the follicular phase (~10 days). In contrast, when pregnancy is established, progesterone levels remain at 50–100 ng/ml for approximately 12 wk and then rise gradually to around 200 ng/ml until just before parturition.
The levels of progesterone in the peripheral plasma of both animals (345W and 457W) that received biparental IVF embryos rose steadily to about 100–120 ng/ml and remained high, typical of progesterone profiles of pregnancy (Fig. 1A). Of the three animals that received parthenogenetic embryos, two animals (469W and 491W) had progesterone profiles resembling those of pregnant marmosets (Fig. 1C), and one recipient (323W) had a progesterone profile typical of a nonpregnant animal (Fig. 1E). Continued high progesterone levels in animals 469W and 491W for 33 days after ovulation would suggest that the CL was maintained, and this was confirmed visually when these recipient marmosets were killed.
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CG Profiles of Recipient Marmosets
Normal levels of CG, which strongly cross-reacts with LH, have also been previously described by Hearn et al. [21]. The natural LH surge and/or administration of exogenous hCG, used to synchronize recipient animals, causes LH/CG to rise sharply around the day of ovulation. If pregnancy is not established, LH/CG levels drop to baseline levels (<40 mIU/ml) and remain low until just before the following ovulation [21]. If pregnancy is established, LH/CG (~50 mIU/ml) can be detected in the peripheral plasma from Day 16–20 and can rise to levels of 1000 mIU/ml by Day 60 of gestation.
Animals that received biparental IVF embryos had LH/CG profiles typical of pregnancy (Fig. 1B). All animals that received parthenogenetic embryos had LH/CG profiles resembling those of nonpregnant animals (Fig. 1, D and F).
Ir-Inhibin Profiles of Recipient Marmosets
Previous studies have shown a significant difference in ir-inhibin levels between conception and nonconception cycles before implantation [22]. During the first 26 days of gestation, animals 345W and 457W (IVF embryo recipients) had total ir-inhibin levels of 2.2 and 1.7 µg x days/ml, respectively. The nonpregnant animal (323W) had a total ir-inhibin over the same 26 days of 1.4 µg x days/ml. However, total ir-inhibin levels of 469W and 491W (parthenogenetic embryo recipients) during the first 26 days of gestation were 2.7 and 2.9 µg x days/ml, respectively. That is, the peripheral plasma ir-inhibin levels of the parthenogenetic embryo recipients were 1.5 times higher than those of the IVF embryo recipients and 2 times higher than those of the nonpregnant animal.
Histology of Uteri after Biparental IVF Embryo Transfer
The in vivo morphology of the biparental IVF embryos was similar to that described previously [13]. Cytotrophoblast displayed extensive but superficial attachment to the luminal surface of the endometrium, covering almost the entire uterine lumen. The chorionic membrane attached to the luminal endometrium and the developing fetus with amniotic sac was clearly visible. The invasion of the syncytiotrophoblast into the stromal tissue of the endometrium could be seen as "finger-like" projections of tissue displaying multinucleated cells typical of syncytium. This cell layer surrounded the blood vessels underlying the apical endometrial epithelium, which had undergone a typical hypertrophy and proliferative response (Fig. 2B). By comparison, the apical endometrium of a nonpregnant marmoset showed no hypertrophy of blood vessels, and the endometrium was of regular appearance (Fig. 2A). The decidual reaction in the marmoset monkey is minor compared to that of other primates [13], so the lack of a decidual reaction in Figure 2B did not necessarily indicate that the animal was not pregnant.
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Histology of Uteri after Parthenogenetic Embryo Transfer
In the two females that displayed biochemical evidence of pregnancy (469W and 491W), there was also histological evidence of implantation, although in both cases the morphology was distinct from that of females pregnant with biparental IVF embryos. In cross sections of the uterus of 491W (Fig. 2D), a plaque of tissue was observed near the apical surface of the endometrium, and presumptive syncytiotrophoblast surrounded blood vessels underlying the epithelium. Syncytial tissue in the stroma, similar to that seen in the normal pregnant animals, was observed. Although remnants of membranes were present on the surface of the endometrium, a definite membrane could not be determined, and there was no discernible fetal tissue. Also present were mitotic bodies, which indicate the continued proliferation of syncytial trophoblast despite the degeneration of the fetal tissue. Cross sections of the uterus of female 469W (Fig. 2C) also exhibited a plaque of putative trophectoderm and a more distinct decidual reaction. Epithelium near the plaque was disorganized, but neither placental membranes nor fetal tissue was observed. Syncytium, which had histology very similar to that seen in normal pregnant animals, was underlying the endometrial epithelium.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Marmoset oocytes underwent four different types of development after activation, similar to those reported for human [24] and mouse [7] oocytes. The majority of ethanol-activated marmoset oocytes underwent immediate cleavage. This may have been due to the age of the oocytes at treatment since aging affects the type of murine development after parthenogenetic activation [25], and meiotic spindles migrate to the center of mouse oocytes during the aging process [26]. Increased immediate cleavage of mouse oocytes was observed after 36 h of in vitro culture [25], and this effect may also have occurred in marmoset oocytes but over a longer time span (3 days in vitro culture). The culture time required to observe this effect may be related to the length of the cell cycle in preimplantation stages: marmoset embryos take four days to reach the 8-cell stage [14], the same length of time taken for mouse embryos to reach the blastocyst stage [27].
Histological analysis of recipient uteri after parthenogenone transfer showed invasion of syncytium into the stromal tissue of the endothelium, which indicated that parthenogenetic marmoset embryos developed and implanted. Postimplantation development of parthenogenetic marmoset fetuses appeared to be minimal, as only remnants of membranous material were present in the uterine lumen 33 days after ovulation. At what stage embryonic loss occurred is difficult to determine purely from the extent of syncytial invasion. The presence of mitotic bodies in the syncytium indicated that this invasion continued after loss of any tissue remaining in the uterine lumen.
In primates, the most important signal yet described for the maternal recognition of pregnancy is CG [28]. Marmosets passively or actively immunized against human CG ß-subunit during the first 6 wk of gestation lose their pregnancies [29]. Therefore, hormone profiles shown here for recipients of parthenogenetic embryos were interesting because both progesterone and ir-inhibin, produced by the CL in marmosets [28], were maintained at levels that would be associated with continued luteotropic support. However, CG, the major luteotropin provided by the embryo, remained at nonpregnant levels in the peripheral plasma. Despite these low levels of CG, parthenogenetic embryos did implant, as determined histologically. Marmoset parthenogenones may have produced enough CG to reach the ovary and cause a luteotropic effect, but CG was not produced in quantities sufficient to be measured in the peripheral circulation. Webley et al. [30] have shown that marmoset luteal cells in vivo are exposed to an unknown luteotropin 2 days after implantation, before CG can be measured in the maternal plasma. These workers propose that the CL may be able to respond to extremely small amounts of CG but do not rule out the possibility that some other luteotropic or anti-luteolytic factor maintains the CL while CG rises to concentrations able to provide luteotropic support [30]. Evidence from inhibin studies in marmosets supports the latter possibility, as there is a significant rise in inhibin production by Day 8 after ovulation in conception cycles compared to nonconception cycles. At this early stage of development, the production of CG by marmoset embryos is undetectable both in vitro and in vivo [22]. It is possible that the embryo produces another factor for maternal recognition at preimplantation stages that causes this significant rise in inhibin production from the CL.
Alternatively, it is possible that parthenogenetic embryos do not produce bioactive CG, as there is some evidence that the gene for the ß-subunit of CG may be imprinted and only expressed from the paternal chromosome [31, 32]. In humans, the ß-subunit of CG is situated on chromosome 19 [33] in an area syntenic to the distal part of mouse chromosome 7 [34], which has some areas known to be imprinted [35]. De Groot et al. [32] showed that the abundance of total ß-subunit of CG in maternal serum is proportional to the number of paternal genomes carried by hydatidiform moles (2 paternal), partial moles (2 paternal, 1 maternal), triploid conceptuses (2 maternal, 1 paternal), and normal conceptuses (1 paternal, 1 maternal). Additionally, the levels of ß-hCG in the serum of human triploid conceptuses with one paternal and two maternal genomes are abnormally low compared to those that carry two paternal genomes [36]. Since parthenogenetic embryos do not carry paternal genomes, an imprinted gene for ß-CG could explain the apparent absence of CG in the maternal plasma.
Implantation of parthenogenetic embryos has been reported in other species [8, 37]. This is the first report of an embryo consisting of purely parthenogenetic cells being capable of implantation in primates, although a human parthenogenetic chimera has been reported [38]. That primate embryos can implant without the participation of a paternal genome highlights the need to minimize levels of parthenogenetic activation associated with human assisted reproductive technologies that result in embryo transfer. Parthenogenetic marmoset embryos provide a useful model system for elucidating the roles of the parental genomes in primate postimplantation development and may facilitate the investigation of the earliest signals for maternal recognition of pregnancy in primates.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Correspondence and current address: Vivienne S. Marshall, Wisconsin Regional Primate Research Center, University of Wisconsin, 1223 Capitol Court, Madison, WI 53715–1299. FAX: 608 263 3524; marshall{at}primate.wisc.edu
3 Current address: Melbourne IVF, East Melbourne, Victoria 3002, Australia.
4 Current address: Department of Molecular Biology and Biotechnology, University of Sheffield, U.K.
Accepted: August 6, 1998.
Received: May 26, 1998.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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