Parthenogenetic activation of mouse oocytes induced by inhibitors of protein synthesis

The mechanisms responsible for the pre-fertilization meiotic block, occurring in the oocyte of most mammals at the second metaphase stage, are unknown. Studies with frog oocytes suggest that Metaphase II-arrest results from the production of a cytostatic factor, as yet unidentified, in the cytoplasm of the secondary oocyte, and this factor disappears following fertilization or artificial activation (Masui & Markert, 1971). When the oocytes of the marine invertebrate Chaetopterus are treated with cycloheximide, meiosis is resumed and the oocytes proceed from Metaphase I to the emission of the second polar body (Zampetti-Bosseler, Huez & Brachet, 1973). Artemia salina eggs remain at the Metaphase I stage of meiosis following sperm penetration when they are removed prematurely from the uterus but if treated on removal with low concentrations of cycloheximide (∼ 10⁻⁷ M) normal development ensues (Fautrez & Fautrez-Firlefyn, 1974). The results presented in this paper indicate that protein synthesis is necessary for the maintenance of the meiotic block at Metaphase II in mouse oocytes since they will complete meiosis and develop parthenogenetically when protein synthesis is inhibited.

Oocytes were released from the oviducts of superovulated 6–8 week old CD1 mice (Charles River, Italy) at various times after the injection of human chorionic gonadotrophin (HCG) (Edwards & Gates, 1959). The oocytes and surrounding cumulus cells were incubated in a mouse embryo culture medium (Whittingham, 1971) supplemented with 10 mM HEPES buffer and the required concentration of cycloheximide (Calbiochem). The oocytes from the right and left oviducts of each female were allotted to different experimental treatments in in order to minimize biological variation. After varying periods of incubation in cycloheximide, the oocytes were washed free of inhibitor and then cultured in the standard embryo culture medium. The total period of incubation including exposure to cycloheximide was always 6 h. All incubations were carried out at 37 °C in 5 ml test-tubes gassed with a mixture of 5 % O₂, 5 % CO₂ and 90 % N₂. Following incubation the oocytes were treated with hyaluronidase and washed in tissue culture medium to remove the cumulus cells. Degenerated oocytes were discarded and the remaining oocytes were fixed in 10 % buffered formalin before they were examined for activation with Nomarski optics at 500 x magn. Oocytes were scored as activated if pronuclear formation had occurred.

Oocytes removed h after the injection of HCG were also treated with varying concentrations of puromycin (Calbiochem) or Actinomycin D (Serva) for 6 h and then examined for signs of activation.

To assay protein synthetic activity in oocytes exposed to cycloheximide, oocytes were cultured for 1 h in a medium containing varying concentrations of cycloheximide and tritiated leucine ([4,5-³H]L-leucine; 40 μCi ml^-1, specific activity 60 Ci mmol^-1, New England Nuclear). After incubation they were washed five times in culture medium containing cold leucine. Samples of 15 oocytes were disrupted by freezing and thawing three times and 100 μg BSA was added to each sample as a carrier protein to facilitate the recovery of the oocyte proteins following precipitation with 10 % cold trichloroacetic acid (TCA). The precipitate was washed three times with 5 % TCA (100 μl/wash), dissolved in 20 μl of 0·8 N-KOH and counted in a toluene based scintillation fluid containing 2 % Biosolv (Beckman). Background values were obtained by counting aliquots of the fluid from the final wash. At each cycloheximide concentration three to four samples of oocytes were prepared and counted.

Oocytes from C57BL and F₁ (C57BL ×; A2G) hybrid females were activated by treatment with 5–10 μg^-1 cycloheximide for 6h and transferred to the oviducts of pseudopregnant recipients either as pronucleate oocytes immediately following incubation in the inhibitor or as 2-cell ova following a further 24 h culture in standard medium. The recipients were either MFI (Olac) or F₁ (C57BL × CBA)hybrid females that were mated with sterile males (carrying the T145H translocation, Lyon & Meredith, 1966). Both stages were transferred on the first day of pseudo-pregnancy, i.e. the day on which the vaginal plug was found and the females were killed and examined for implantation sites 6 days later (the seventh day of pseudopregnancy). All implantation sites were fixed and prepared for histological examination.

Recently ovulated mouse oocytes at the Metaphase II stage (⁠h post-HCG) undergo parthenogenetic activation as indicated by the formation of pronuclei when incubated for 6 h in the presence of cycloheximide (Fig. 1). The results of two experiments are combined in the figure; an overall total of 934 oocytes were observed (53–220 oocytes at each concentration). The percentage of activated oocytes increased with the concentration of cycloheximide. Activation was also induced by treatment of oocytes for 6 h with varying concentrations of puro-mycin (Table 1) but the activation response declined with exposure to concentrations higher than 10 μg ml^-1. Examples of the four major types of parthenogenones (Tarkowski, 1975) were produced (Fig. 2) although the group with one pronucleus and second polar body was predominant, e.g. out of 74 oocytes activated in the presence of 10 μg ml^-1 cycloheximide, 66 had one pronucleus and the second polar body, two had immediately cleaved (Braden & Austin, 1954), two had one pronucleus and no second polar body and four had two pronuclei and no second polar body. The morphological appearance of the activated oocytes was normal, the pronuclei were well formed and contained one or more nucleoli. Incubation of oocytes with Actinomycin D (0·05 and 0·1 μg ml^-1) failed to initiate activation above control levels.

In the next series of experiments Metaphase II oocytes at h post-HCG were treated with various concentrations of cycloheximide for 1 h, followed by culture in a standard medium for a further 5 h. The results of two experiments are combined in Fig. 3; an overall total of 1099 oocytes were examined (79–190 oocytes at each concentration). The dose-response curve was found to be more complex than the activation response observed after prolonged incubation in cycloheximide (Fig. 1); a peak of activation was obtained at 1 μgml^-1 cycloheximide followed by a second rise in activation at concentrations greater than 3 μgml-¹. When the response is compared with the incorporation of [³H]leucine in the presence of inhibitor,oocytes are activated by concentrations of cycloheximide that depress protein synthesis by more than 70 %. No significant differences were found in the rate of decay of the proteins synthesized in the presence of various concentrations of cycloheximide (0·01–10 μg ml^-1) and it was also found that oocytes were incorporating levels of labelled leucine similar to the untreated controls within 20 min of removal from 10 μg ml^-1 cycloheximide (unpublished observations).

A qualitatively similar response was obtained when oocytes were treated with a high concentration of cycloheximide (10 μg ml^-1) for varying periods of time (Fig. 4a-c) instead of treating oocytes for a fixed period of time in varying concentrations of the inhibitor. The results of 2–3 experiments are combined in the figure; overall totals of 554, 890 and 950 oocytes were examined at 14, and h post-HCG respectively (48–190 oocytes at each exposure time). The initial activation peak occurs after a period of exposure to the inhibitor which becomes progressively shorter with the increase in post-ovulatory age of the oocyte (in CD1 females ovulation is completed by approx. 12 h after the injection of HCG, unpublished observations). If lower concentrations of inhibitor are used (0·3 μg mb¹) the initial peak of activation is not discernible (Fig. 5). Three experiments are combined in the figure; an overall total of 1243 oocytes were examined (86–220 oocytes at each exposure time). The response reaches a maximum after 3–4 h exposure after which there is a slight decline.

Oocytes activated with cycloheximide were capable of further development following transfer to the oviducts of pseudopregnant recipients (Table 2). Although the number of oocytes transferred was comparatively small, the proportion of activated oocytes developing to the stage capable of initiating a decidual response was similar to the development of oocytes activated in other ways (see reviews by Graham, 1974; Tarkowski, 1975). The decidual cell reaction appeared normal but giant cells and embryonic tissue were only discernible in a small proportion of the implantation sites. In the few instances where the implantation sites contained embryos, embryonic development was retarded by approximately 24 h. Haploid embryos originating from inbred rather than hybrid mice did not appear to have a greater development potential as suggested previously (Kaufman, Huberman & Sachs, 1975). The larger number of implantation sites resulting from the transfer of activated oocytes after culture to the 2-cell stage (13/21–62 %) may be due to the extra time the embryos spent in utero before implantation, for haploid blastocysts have significantly lower cell numbers when compared with diploid controls (fertilized) of similar developmental age due to a slower cleavage rate after the 4-cell stage (Kaufman & Sachs, 1976).

Our results show that the Metaphase II oocyte of the mouse synthesizes protein factor(s) which are necessary for the maintenance of the meiotic block. A similar phenomenon may be responsible for the meiotic blocks occurring in the oocytes of other classes of animals (Masui & Markert, 1971; Zampetti-Bosseler et al. 1973; Fautrez & Fautrez-Firlefyn, 1974). The formation of pronuclei in the presence of cycloheximide or puromycin indicates that this process is not dependent upon the synthesis of new proteins and the inability of Actinomycin D to activate the oocytes suggests that the synthesis of the inhibitory protein(s) is not controlled at the transcriptional level. The treatment of oocytes with cycloheximide for 6 h does not impair their ability to develop at least to the stage of implantation.

‘Activating’ effects of various types can be induced in other repressed cellular systems by treatment with inhibitors of protein synthesis. In serum-starved and density-inhibited human fibroblasts cycloheximide causes an increase in putrescin transport which is a characteristic response associated with the initiation of cell proliferation (Pohjanpelto, 1976). Uridine uptake is increased in serum-deprived mouse fibroblasts treated with cycloheximide (Hershko,Mamont, Shields & Tomkins, 1971) and RNA synthesis is stimulated in aminoacid-deprived HeLa cells treated with cycloheximide (Smulson & Thomas, 1969). These activating effects are usually attributed to the inhibition of the synthesis of rapidly metabolising repressor proteins.

The activation of the oocytes may not be due simply to the removal of ‘blocking’ agents since with short exposures to cycloheximide the activation response does not increase progressively with increasing concentrations of inhibitor. From our data, it would appear that the meiotic block and subsequent activation are regulated not only by ‘blocking’ protein factor(s) but also by other protein(s) having opposite effects and different rates of synthesis or decay. Thus, the response obtained with a low concentration of cycloheximide (Fig. 5) might indicate that under these conditions the intracellular concentration of blocking factors is affected more than the factors necessary for activation. The ratio between the 2 factors probably varies during the natural ageing of the oocyte during the postovulatory period, since there is an increase in spontaneous activation with age (Austin, 1961 ; Longo, 1974), and an earlier occurrence of the initial activation peak in the presence of inhibitor with advancing postovulatory age (Fig. 4 a-c). Furthermore, the fertilizability of ovulated mouse eggs also varies with time in a way parallel to activation with cycloheximide: a dramatic increase occurs between 13 and 15 h post-HCG, and maximal fertilizability is reached at 17 h (Iwamatsu & Chang, 1971).

We thank Mr M. Coletta and Mrs J. Keogh for technical assistance. This work was supported by the World Health Organization, the Ford Foundation, the Consiglio Nazionale delle Ricerche (Biology of Reproduction Grant no. 7600300.85) and NATO.