1 Tsukuba Primate Center for Medical Science, National Institute of Infectious Diseases, Hachimandai-1, Tsukuba, Ibaraki, 3050843, 2 Bioresource Engineering Division, Bioresource Center, RIKEN Tsukuba Institute, Ibaraki, 3 Department of Veterinary Science, National Institute of Infectious Diseases, Tokyo, 4 National Institute for Minamata Disease, Kumamoto and 5 Tokyo University of Agriculture, Hokkaido, Japan
6 To whom correspondence should be addressed. e-mail: sankai{at}nih.go.jp
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Abstract |
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Key words: cynomolgus monkey/embryo transfer/oocyte activation/ROSI/round spermatid
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Introduction |
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We have previously confirmed the technical basis for embryo culture, embryo transfer (Sankai et al., 1994; Sankai, 2000
), and ICSI (Ogonuki et al., 1998
) in non-human primates, especially the cynomolgus monkey (Macaca fascicularis). By using the ICSI and ROSI techniques, we found that not only mature sperm but also round spermatids from cynomolgus monkeys can activate mouse oocytes by inducing repetitive intracellular Ca2+ increases (Ca2+ oscillations) (Ogonuki et al., 2001
). In this regard, cynomolgus monkey round spermatids are very similar to human round spermatids (Yazawa et al., 2000
). Although, in the rhesus monkey, an infant was born from embryos after injection of elongated spermatids (Hewitson et al., 2002
), the round spermatids of this species do not have oocyte-activating capacity (Hewitson et al., 2000
). Therefore, the cynomolgus monkey may be a potential experimental model for a better understanding of human ROSI. In this study, we injected round spermatids from cynomolgus monkeys into homologous mature oocytes and examined their ability to support embryo development in vitro and in vivo.
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Materials and methods |
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On the day of the experiment, a cryotube was placed in a water bath at room temperature. When the suspension began to thaw, 1 ml of GL-PBS was added. The mixture was then poured into a 15 ml tube containing 5 ml of GL-PBS. After complete thawing, the suspension was pipetted gently and washed twice by centrifugation using GL-PBS. To prevent the cells from losing their viability, the cell suspension was kept at 4°C until just before use.
Preparation of oocytes
Eight sexually mature female cynomolgus monkeys, whose menstrual cycles were confirmed to be normal (2832 days) by examining vaginal bleeding (Honjo et al., 1984) for at least three cycles, were used for oocyte collection. Females received multiple i.m. injections (two times per day; 1214 times total) of hMG (Pergonal; Teikoku Hormone, Japan). hCG (Profasi; Ares Serono, USA) was injected i.m. on the morning after the last hMG injection. In four of the eight females, to inhibit spontaneous release of LH or FSH from the pituitary gland, GnRH agonist (Leuplin; Takeda, Japan) was injected on the first day of menses; 2 weeks later, the four females received the first of multiple injections of hMG. Between 28 and 43 h after the hCG injection, all females were anaesthetized with a combination of 10 mg of ketamine hydrochloride per kg of body mass (Ketalar) and 1 mg of xylazine hydrochloride per kg of body weight (Seraktarl; Bayer, Germany). Ovaries were exposed through an abdominal incision (Figure 1a), and the contents of enlarged follicles (15 mm in diameter) were aspirated through a 25 gauge needle connected to a 2.5 ml syringe. The collected follicular contents were diluted immediately with HEPES-TYH medium (Toyoda et al., 1971
) containing 2.5 IU/ml of heparin (Novo Nordisk, Denmark), and the oocytes were rinsed with TYH medium consisting of 119.37 mmol/l NaCl, 4.78 mmol/l KCl, 12.6 mmol/l CaCl22H2O, 1.19 mmol/l MgSO47H2O, 1.19 mmol/l KH2PO4, 25.07 mmol/l NaHCO3, 5.56 mmol/l glucose, 1.0 mmol/l pyruvic acid (sodium salt), 5 mg/ml BSA, penicillin-G (sodium salt), and streptomycin sulphate. HEPES-TYH was made by substituting HEPES for 20 mmol/l NaHCO3. Heparin was added to the HEPES-TYH medium to prevent coagulation of contaminating blood.
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Microinjection of round spermatids
Microinjection was done using a micromanipulation system equipped with a piezo-micropipette-driving unit (Prime Tech, Japan) combined with a Nomarski interference-contrast microscope (Nikon Optical Co., Japan) (Figure 2). This procedure was reported previously (Kimura and Yanagimachi, 1995a,b; Ogonuki et al., 1998
). The cover of a plastic dish (Falcon no. 1006; Becton Dickinson, USA) was used as the microinjection chamber. We remodelled the covers to use a Nomarski interference-contrast microscope by hollowing out a 2 cm diameter section and then placing over this section a glass coverslip with silicon-based adhesive around its circumference. Several small drops (25 µl), each containing HEPES-TYH (for oocytes) and 12% PVP in HEPES-TYH, were placed on the coverslip. In one of these drops, spermatogenic cells were suspended to a final PVP concentration of 6%. The drops were then covered with silicone oil (Aldrich Chemical Co., USA). The monkey round spermatids were
1012 µm in diameter (Figure 2a) and were thus easily distinguished from primary spermatocytes (1720 µm in diameter). A round spermatid was drawn into the injection pipette (Figure 2b-1). After the plasma membrane was broken by pipetting, the nucleus with a small amount of the cytoplasm of a round spermatid was injected into the ooplasm (Figure 2b-24). Oocytes injected with round spermatids were transferred into TYH medium, covered with silicone oil, and then kept at 38°C in a humidified 5% CO2 and 5% O2 atmosphere with N2 gas.
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Embryos obtained by ROSI were co-cultured with buffalo rat liver cells (Zhang et al., 1994) at 23x104 cells per well of a dish for IVF (Falcon no. 353653; Becton Dickinson) in CMRL-1066 medium supplemented with 10% FCS. The progression of embryo growth was examined daily by using a Hoffman modulation contrast microscope (Leica; Heerbrugg, Switzerland).
Embryo transfer
Five females were used for tubal embryo transfer. Ovulation in the menstrual cycle of the females was estimated by measuring the serum concentrations of estradiol using a commercially available enzyme immunoassay kit for human hormones (IMx system; Dainabot Co., Ltd) (Ogonuki et al., 1997). Blood samples were taken in the morning, and sera separated by centrifuge were frozen until the measurements were taken. The day of ovulation was estimated as the day that the estradiol concentration significantly decreased. Approximately 4667 h post-ROSI, embryos at the 616-cell stage were selected for tubal transfer. These criteria were based on our previous experience. Embryos were picked from the culture dishes into a micro-glass capillary containing 35 µl of culture medium. Synchronizing the developmental stage of each embryo with the menstrual cycle of the recipient, tubal embryo transfer was done 23 days after ovulation. Females were anaesthetized with a combination of ketamine hydrochloride and xylazine hydrochloride. Fallopian tubes and ovaries were exposed through an abdominal incision, and the micro-glass capillary containing the embryos was introduced through the fimbriated end of the Fallopian tube; one embryo per tube was transferred into the mid-ampullary portion of the oviduct.
At 28 days post-ROSI, the presence of a gestational sac and the heartbeat of a fetus were confirmed by ultrasonography. Subsequently, the growth of the fetus was examined at 56 and 84 days post-ROSI.
Statistical analysis
Results were evaluated using Fishers exact test; P < 0.05 was considered significant.
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Results |
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Discussion |
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The first fertilization using round spermatids was reported in hamsters (Ogura and Yanagimachi, 1993). It is now generally accepted that mouse round spermatids can support full-term embryo development following ROSI with an efficiency comparable with that of mature sperm (Ogura et al., 1994
; 2001; Kimura and Yanagimachi, 1995b
). ROSI in other animals, including rabbit (Sofikitis et al., 1994
) and rat (Hirabayashi et al., 2002
), has also been reported to result in the birth of normal offspring. However, the use of round spermatids as substitute gametes in humans has been controversial. Some infertility clinics have achieved successful birth (Fishel et al., 1995
; 1996; Tesarik et al., 1995
; 1996; Antinori et al., 1997
), but many clinics have experienced very low fertilization rates following ROSI (Yamanaka et al., 1997
; Levran et al., 2000
; Vicdan et al., 2001
). This discrepancy between humans and other animals in round spermatid conceptions may be attributable either to species-specific differences in round spermatids or to circumstances in which round spermatids are collected from infertile humans with spermiogenic failure. According to Yazawa et al. (2000
), human round spermatids essentially have oocyte-activating capacity while inducing intracellular Ca2+ oscillations, as demonstrated by the assay system using mouse mature oocytes. The present study indicates that, once oocytes of cynomolgus monkeys are successfully activated by round spermatids, many of them undergo normal preimplantation development, and some continue their development at least until mid-gestation. Thus, it is very probable that the poor outcome in human ROSI is caused by the impaired oocyte-fertilizing ability, or, more specifically, the oocyte-activating capacity, of round spermatids collected from individual patients. However, this assumption is incompatible with a finding by Hewitson et al. (2000
), who reported an extremely low efficiency of ROSI in the rhesus monkey (Macaca mulatta) whereas injection of elongated spermatids was successful in the same species. Because the stage at which spermatogenic cells acquire the oocyte-activating capacity is species-dependent (Fishel et al., 1997
; Sofikitis et al., 1997
; Yazawa et al., 2000
; Ogonuki et al., 2001
), the detailed examination of rhesus monkey round spermatids for oocyte-activating capacity is important for resolving the discrepancy between the two macaque species.
We performed ROSI using oocytes that matured in vitro as well as in vivo. No significant differences in percentages of pronucleus formation, cleavage, morula, and blastocyst formation were noted between the two groups. However, only a small percentage of oocytes matured from the germinal vesicle stage during this culture period, which varied from 9 to 30 h. The most serious problem we encountered was a fluctuation in the quality of oocytes, which varied greatly among different donor females. At present, it is very difficult to obtain monkey oocytes of good quality. Many oocytes at the time of collection are either overmatured or are too immature (at the germinal vesicle stage), and only a few immature oocytes undergo maturation in vitro. More reliable methods for ovarian stimulation or for in-vitro maturation are needed to obtain oocytes of good quality that will ensure consistent experimental outcomes. Such technical improvements may further increase the rates of normal fertilization and embryo development following ROSI and will make ROSI a common artificially assisted reproduction technique in non-human primates.
In cynomolgus monkeys, the fertilization rate by conventional IVF (assessed by pronucleus formation) is 57% (Sankai et al., 1994), whereas the fertilization rate by ROSI is 69% (the present study). This is because sperm with a high motility are necessary for successful IVF, while ICSI and ROSI do not require motility of male germ cells. It is also advantageous that cells which had been frozenthawed by a simple method can also be used for ICSI into oocytes. Thus, in cynomolgus monkeys we obtained preimplantation embryos more easily by ROSI than by IVF.
This does not necessarily mean that offspring can be obtained easily by ROSI, because ours is the first known case of pregnancy of a cynomolgus monkey by ROSI; there have been no other reports of pregnancy following ROSI in a non-human primate. Sofikitis et al. (1996) reported that electrical stimulation of rabbit oocytes before round spermatid nuclei injection (ROSNI) had beneficial effects on oocyte activation, fertilization and development. Yazawa et al. (2000
) reported that rabbit round spermatids activated mouse oocytes but could not induce Ca2+ oscillation; normal Ca2+ oscillation patterns were seen in only three of nine oocytes. In contrast, our earlier report (Ogonuki et al., 2001
) showed that cynomolgus monkey round spermatids could activate mouse oocytes (57/60; 95%) with normal Ca2+ oscillation (10/14; 71%). From these two reports, we conclude that monkey round spermatids have a higher concentration of oocyte-activating factor or have a more mature form than rabbit round spermatids. If we use spermatids that have little or no oocyte-activating factor, such as those from mice or rabbits, electrical stimulation has a beneficial effect. We cannot predict whether electrical stimulation would produce good results for ICSI of cynomolgus monkey spermatids that have already matured and acquired oocyte-activating capacity.
Thus, although the ROSI technique has been established in many animals, the species in which ROSI is successful need chemical or electrical stimulation to activate the oocyte. Cynomolgus monkeys are the only species other than humans in which the round spermatids like human round spermatids have oocyte-activating factor. We must have an animal model that is similar to the infertile human with spermiogenic failure. We therefore believe that the cynomolgus monkey is a suitable model for humans.
In the colony of cynomolgus monkeys at our institute, spontaneous abortion rarely occurs, and at present we do not know the reason for the spontaneous abortion of the ROSI fetus. It may have been due to either technical or maternal factors. Nevertheless, in this study, we succeeded in the first post-implantation development of a ROSI fetus that lasted to day 103 of the 156 day pregnancy period of the cynomolgus monkey. Because the safety and effectiveness of round spermatid conception in humans are still far from certain because of the lack of substantial scientific evidence (Aslam et al., 1998; Sofikitis et al., 1998b
), cynomolgus monkeys are valuable experimental models for assessing the application of this technique to treatment for human infertility. In the future, the ROSI technique will also be applicable to rescue of endangered wild animals in which IVF and ICSI have never been successful.
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Acknowledgements |
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References |
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Submitted on April 12, 2002; resubmitted on November 4, 2002; accepted on January 21, 2003.