The effect of balanced gene dosage changes on the timing of cavitation and on the timing of appearance of a stage-specific embryonic cell surface antigen was studied in preimplantation mouse embryos. Gene dosage was increased by creating tetraploid embryos at the 4-cell stage, either by blastomere fusion with polyethylene glycol (PEG) or by incubation in cytochalasin B (cytB) to block cell division. Removal of the zona pellucida with Pronase from diploid embryos caused a 7 h delay in cavitation. Further manipulations, either with PEG or cytB to induce tetraploidy, did not produce a statistically significant additional delay in cavitation timing. Likewise, PEG-induced tetraploidy did not affect the timing of appearance or disappearance of the embryonic cell surface antigen as compared with diploid control embryos. In analysing the metabolic effects of tetraploidy, we found that in tetraploid embryos with cell number equivalent to intact diploid embryos, MDH activity did not double with the doubling of the genome, being only 50 % greater than diploid levels in cytB-induced tetraploid embryos and only 20 % greater than diploid levels in PEG-induced tetraploid embryos. However, in tetraploid embryos with one-half normal cell number, enzyme activity was equal to that in whole diploid embryos, suggesting that in such embryos, MDH activity increased in parallel with increases in gene dosage. Further studies showed that levels of RNA synthesis in PEG-induced tetraploid embryos also did not increase in parallel with the doubling of the genome. Rather, these results suggested that in tetraploid embryos, compensation was made for at least part of the excess genetic material.

Although preimplantation development in the mouse is defined by numerous morphological, biochemical, and physiological changes, there still remains the problem of how their timing and coordination are controlled. This study was initiated to investigate the control of the timing of formation of the blastocoele (i.e. cavitation).

Recent hypotheses implicate the nucleus in the timing of preimplantation development. The maternal uterine environment cannot be providing any critical cues, since development in vitro proceeds normally and at very nearly the rate observed in vivo. Likewise, cytokinesis per se must not be critical in the control of timing, since the number of cells and cell divisions can be altered without affecting the timing of blastocyst formation (Smith & McLaren, 1977). However, neither the number of nuclear divisions (i.e. DNA replications) nor changes in the ratio of nuclear-to-cytoplasmic volume have been excluded from consideration as possible repositories of the putative cue for cavitation (Smith & McLaren, 1977; Alexandre, 1979; Braude, 1979; Surani, Barton & Burling, 1980).

The embryonic genome is known to be active during preimplantation development, as shown by the detection of mRNA synthesis (Knowland & Graham, 1972) and the expression of paternal isozymes (Chapman, Whitten & Ruddle, 1971; Wudl & Chapman, 1976). Also, the inhibitor of mRNA synthesis, α-amanitin, has been used to show that blastocyst formation is dependent on new mRNA synthesis (Braude, 1979). Since the genome is clearly functioning in early development, it seems reasonable to study the nature of the ‘developmental clock’ by determining whether nuclear changes will affect the timing of cavitation.

In this report, balanced increases in gene dosage to a tetraploid level were used to determine whether they would alter the timing of cavitation. We produced tetraploid embryos with a recently developed method to fuse embryonic blastomeres using polyethylene glycol (PEG; Eglitis, 1980), as well as by the standard procedure using cytochalasin B (cytB; Snow, 1973). The interaction of tetraploidy with cell number changes and its effect on developmental timing were analysed by monitoring cavitation rates and the expression of a stage-dependent embryonic cell surface antigen. Additional biochemical studies were done to monitor the effect of tetraploidy on embryonic metabolism.

Embryos at the 2-cell stage (47 ∼ 49h after hCG injection) were flushed from the oviducts of superovulated, random-bred DUB: (ICR) female mice with bicarbonate-free modified Hank’s balanced salts solution (BSS; Spindle & Goldstein, 1975). Embryos were grown in modified (Spindle & Goldstein, 1975) standard egg culture medium (Biggers, Whitten & Whittingham 1971) under paraffin oil (Fisher) and incubated at 37°C in a humidified atmosphere of 5%CO2 in air.

Depending upon the particular experiment, zonae pellucidae were removed either from diploid 2-cell embryos before experimental manipulations (for blastomere fusion) or from tetraploid 2-cell embryos after experimental manipulations (for cytB). Zonae pellucidae were removed by a 5 min incubation at 37 °C in 0·5 % protease (Pronase, Sigma; Mintz, 1962).

Tetraploid embryos were obtained by two methods: blastomere fusion (Eglitis, 1980), or, inhibition of cell division with cytB (Snow, 1973). Briefly, fused blastomeres were produced by disaggregating 4-cell-stage embryos, aggregating individual blastomeres in pairs with phytohaemagglutinin (PHA, Mintz, Gearhart & Guyment, 1973), and treating aggregated pairs for 2 min with 45 % (w/v) PEG (Sigma, MW 1000; see Eglitis, 1980; Fig. 1). Alternatively, tetraploid embryos were produced by incubating 2-cell embryos for 13 h (from 49 -62 h post-hCG) in standard egg-culture medium containing 10 μg cytB (CalBiochem) per ml of medium.

Fig. 1.

A diagram illustrating the general features of the blastomere fusion protocol. For a full description, see Eglitis (1980).

Fig. 1.

A diagram illustrating the general features of the blastomere fusion protocol. For a full description, see Eglitis (1980).

To investigate the affect on developmental timing of the interaction of tetraploidy and altered cell number, both fusion and cytB-induced tetraploid blastomeres were aggregated with PHA in various cell number combinations. Fused blastomere tetraploid embryos (PEG-4N) were followed after aggregation to restore cell number to that of equivalent (i.e. 4-cell) intact diploid embryos. Alternatively, development of such tetraploid embryos with only half (; PEG-4N) or one-quarter ( PEG-4N) the cell number of intact embryos was also monitored. CytB-induced tetraploid embryos were followed either as ‘half’-embryos with ( w/Z-cytB-4N) or without ( ZF-cytB-4N) zonae pellucidae, or, after zona removal, as aggregated to restore cell number to that of equivalent intact diploid embryos (cytB-4N).

Controls consisted of intact diploid embryos with (w/Z-2N) or without (ZF-2N) zonae pellucidae, as well as diploid embryos disaggregated at the 4-cell stage to ( ZF-2N) or ( ZF-2N) diploid embryos. Additional controls included blastomeres which did not fuse after PEG treatment, either reaggregated to restore cell number to that equivalent to intact embryos (PEG-2N) or grown without reaggregation ( PEG-2N).

Two parameters were used to time development of the embryos: (1) timing of blastocoele formation, and (2) timing of expression of a stage-specific embryonic cell surface antigen.

To time progress towards cavitation, embryos were checked at intervals through an inverted phase-contrast microscope. Embryos were scored as cavitating when an intraembryonic cavity became visible. The timing of cavitation was compared between different classes of embryos by comparing their of cavitation (i.e. the time at which 50 % of the final maximum proportion of cavitation embryos was reached). To determine the the linear regression was taken over the times at which the proportion of cavitated embryos was uniformly increasing. Then, the was calculated by finding the time at which one-half the final maximum number of embryos had cavitated.

The timing of expression of the embryonic cell surface antigen was monitored by indirect immunofluorescence (IIF). The particular antigen assayed was one detected by an antiserum raised to 8-cell-stage embryos in a male New Zealand white rabbit. The cell surface expression of this antigen (or antigens) is very ‘stage-specific’ in that it is restricted to the period immediately preceding and following the onset of cavitation (around 94–06 h post-hCG). If necessary, zonae pellucidae were mechanically removed immediately before the embryos were observed through a Zeiss phase-contrast microscope equipped with epifluorescence. Negative controls consisted of embryos incubated in the antiembryo antiserum at times when the antigen was not expressed, as well as embryos incubated in pre-immune (normal) rabbit antiserum.

Two parameters were measured to assess the metabolic levels of tetraploid embryos. The first was the determination of the level of activity of the constitutive enzyme, malate dehydrogenase (MDH). The level of MDH activity was determined in diploid and tetraploid morulae (90 h post-hCG) and blastocysts (120 h post-hCG) spectrophotometrically from the rate of reduction of NAD (Brinster, 1966). Enzyme activity in moles NAD reduced/h/embryo was calculated using the formula:
formula
In the absence of substrate, embryos reduced only an insignificant amount of NAD.

The second measure of embryonic metabolism was the determination of the level of RNA synthesis per blastomere. This was accomplished autoradio-graphically by quantitating incorporation of [3H]uridine into nuclear RNA. Late morulae, (92–93 h post-hCG) were incubated for 3 h in standard egg culture medium containing 0·5 or 0·05 μCi/ml of [6-3H]uridine (specific activity 22-4Ci/mM, New England Nuclear; concentration 2·23 × 10−5 or 2·33 × 10−6 mM, respectively). Labelled embryos were rinsed thoroughly in BSS containing 2·33 × 10−2 mM cold uridine. After labelling, embryos were fixed onto glass slides by a modification (Epstein, Smith, Travis & Tucker, 1978) of Tarkowski’s (1966) air-drying method. The slides were dried overnight in a dessicator at 4 °C and then dipped in NTB-2 emulsion (Kodak); coated slides were allowed to dry in safe boxes. Embryos labelled in 0·5 μCi [3H]uridine/ml were developed 18 h after dipping, while embryos labelled in 0·05 μCi [3H]uridine/ml were developed 104 h after dipping. Developed slides were counter-stained with methyl green/ pyronin and coverslipped. Grains per nucleus were determined by viewing under oil immersion. Only embryos with cell numbers between 8 and 16 were used to quantitate RNA synthesis so as to minimize cleavage stage-dependent variability in RNA synthesis, and so as to maximize the comparability of experimental embryos with control embryos. Correction for background levels of radioactivity were made by counting the number of grains per nucleus for embryos that were fixed without being incubated in [3H]uridine.

Timing of blastocyst formation

Intact diploid embryos with zonae pellucidae (w/Z-2N) had a for cavitation of 97 h post-hCG, whereas, zona-free diploid embryos (ZF-2N) cavitated about 7 h later (see Table 1, Fig. 2ac). In fact, after the delay in cavitation accompanying removal of the zona pellucida, further manipulations of the embryos, whether to reduce cell number, induce tetraploidy, or both, did not result in a statistically significant change in the timing of cavitation. Regardless of the means used to induce tetraploidy and regardless of whether or not cell number was restored to normal, all classes of tetraploid embryos cavitated at the same time as did ZF-2N embryos (i.e. about 104 h post-hCG; see Fig. 3). In addition, in the case of PEG-treated embryos which did not fuse (i.e. PEG-2N), such embryos also had a for cavitation of around 104 h post-hCG (data not shown).

Table 1.

Timing of cavitation, t12 = h post-hCG

Timing of cavitation, t12 = h post-hCG
Timing of cavitation, t12 = h post-hCG
Fig. 2.

Timing of cavitation. Embryos were monitored by phase contrast microscopy for the presence or absence of a blastocoele.

formula

t12 (time at which 12 the final maximum number of embryos had cavitated) was determined by calculating the linear regression for the times at which the number of cavitated embryos was increasing, then calculating t using the value of 12 ‘% cavitated’ that was reached at the end of the culture period. Arrow on graphs point to the t12 (a) PEG Tetraploids: ∘ - - ∘, with cell number restored to whole control levels (quartets); □- - -□ with cell number equivalent to 12 controls (doublets); ⋆ - - -, ⋆ with cell number equivalent to 14 controls (singlets). (b) Cytochalasin B Tetraploids : ⊙ - - ⊙, with cell number restored to whole control levels (restored-whole) ; ⊡ - - - ⊡, with cell number equivalent to 12 controls (half), (c) Control Diploids: ⭑ — ⭑, whole diploid embryos with intact zonae pellucidae (with zonae); • - - •, while diploid embryos with zonae pellucidae removed at the 2-cell stage with 0·5 % Pronase (zona-free); ◼- - -◼ zona-free diploid embryos with 12 normal cell number (half); ⋆ - - - ⋆, zona-free diploid embryos with 14 normal cell number (quarter).

Fig. 2.

Timing of cavitation. Embryos were monitored by phase contrast microscopy for the presence or absence of a blastocoele.

formula

t12 (time at which 12 the final maximum number of embryos had cavitated) was determined by calculating the linear regression for the times at which the number of cavitated embryos was increasing, then calculating t using the value of 12 ‘% cavitated’ that was reached at the end of the culture period. Arrow on graphs point to the t12 (a) PEG Tetraploids: ∘ - - ∘, with cell number restored to whole control levels (quartets); □- - -□ with cell number equivalent to 12 controls (doublets); ⋆ - - -, ⋆ with cell number equivalent to 14 controls (singlets). (b) Cytochalasin B Tetraploids : ⊙ - - ⊙, with cell number restored to whole control levels (restored-whole) ; ⊡ - - - ⊡, with cell number equivalent to 12 controls (half), (c) Control Diploids: ⭑ — ⭑, whole diploid embryos with intact zonae pellucidae (with zonae); • - - •, while diploid embryos with zonae pellucidae removed at the 2-cell stage with 0·5 % Pronase (zona-free); ◼- - -◼ zona-free diploid embryos with 12 normal cell number (half); ⋆ - - - ⋆, zona-free diploid embryos with 14 normal cell number (quarter).

Fig. 3.

The appearance, development and ploidy of fused-blastomere tetraploid embryos (ac) The appearance of control, ZF-2N embryos, including a representative chromosome spread.

(df) The appearance of experimental, PEG-4N embryos.

(a, d) The appearance of diploid and tetraploid embryos at the end of the experimental manipulations (about 60 hpost-hCG injection). The tetraploid embryo (d) is composed of four fused blastomeres combined by PHA aggregation.

(b, e) Diploid and tetraploid blastocysts about 140 h post-hCG. Note the prominent inner cell mass in both diploid and tetraploid embryos. The tetraploid blastocyst developed from a restored four cell form as illustrated in (d) and was photographed 83 h after PEG-induced fusion.

(c,f) Metaphase plates obtained from diploid and tetraploid-embryos. Blastocysts were fixed 139 h post-hCG, after a 3h preincubation in 0·2 μg/ml colcemid to maximize cells in metaphase, according to a modification (Epstein et al. 1978) of Tarkowski’s (1966) method. Magnification – bar in e applies to a, b, d and e and represents 50 μm. Bars in c and f represent 1 μm.

Fig. 3.

The appearance, development and ploidy of fused-blastomere tetraploid embryos (ac) The appearance of control, ZF-2N embryos, including a representative chromosome spread.

(df) The appearance of experimental, PEG-4N embryos.

(a, d) The appearance of diploid and tetraploid embryos at the end of the experimental manipulations (about 60 hpost-hCG injection). The tetraploid embryo (d) is composed of four fused blastomeres combined by PHA aggregation.

(b, e) Diploid and tetraploid blastocysts about 140 h post-hCG. Note the prominent inner cell mass in both diploid and tetraploid embryos. The tetraploid blastocyst developed from a restored four cell form as illustrated in (d) and was photographed 83 h after PEG-induced fusion.

(c,f) Metaphase plates obtained from diploid and tetraploid-embryos. Blastocysts were fixed 139 h post-hCG, after a 3h preincubation in 0·2 μg/ml colcemid to maximize cells in metaphase, according to a modification (Epstein et al. 1978) of Tarkowski’s (1966) method. Magnification – bar in e applies to a, b, d and e and represents 50 μm. Bars in c and f represent 1 μm.

Timing of antigen appearance

At 90 h post-hCG, none of the embryos expressed any antigen on their cell surfaces, as detected by IIF (Fig. 4). However, at both 94 and 101 h post-hCG, PEG-4N, w/Z-2N and ZF-2N embryos expressed this embryonic antigen on their cell surfaces. The antigen was no longer expressed on the trophectoderm of blastocysts tested by IIF 115 h post-hCG. Thus, antigen expression on tetraploid embryos was neither accelerated delayed, nor prolonged when compared with the controls.

Fig. 4.

The expression of a stage-specific embryonic cell surface antigen on diploid and tetraploid morulae.

(ad) Control diploid embryos, grown in culture from the 2-cell stage with intact zonae pellucidae. Zonas mechanically removed for IIF.

(eh) PEG-4N embryos.

(a, e) 90 h post-hCG, PEG-4N embryos 31 h after PEG-induced fusion.

(b,f) 94 h post-hCG, PEG-4N embryos 35 h after PEG-induced fusion.

(c, g) 101 h post-hCG, PEG-4N embryos 42 h after PEG-induced fusion.

(d, h) 115 h post-hCG, PEG-4N embryos 56 h after PEG-induced fusion. All photographs taken with 1·5 min exposures of Kodak Tri-X film. Prints all exposed for 35 sec. In h, bright patches of fluorescence are caused by blastomeres killed during HF manipulations which filled with fluorescent antibody. Magnification -bar in h represents 50 μm.

Fig. 4.

The expression of a stage-specific embryonic cell surface antigen on diploid and tetraploid morulae.

(ad) Control diploid embryos, grown in culture from the 2-cell stage with intact zonae pellucidae. Zonas mechanically removed for IIF.

(eh) PEG-4N embryos.

(a, e) 90 h post-hCG, PEG-4N embryos 31 h after PEG-induced fusion.

(b,f) 94 h post-hCG, PEG-4N embryos 35 h after PEG-induced fusion.

(c, g) 101 h post-hCG, PEG-4N embryos 42 h after PEG-induced fusion.

(d, h) 115 h post-hCG, PEG-4N embryos 56 h after PEG-induced fusion. All photographs taken with 1·5 min exposures of Kodak Tri-X film. Prints all exposed for 35 sec. In h, bright patches of fluorescence are caused by blastomeres killed during HF manipulations which filled with fluorescent antibody. Magnification -bar in h represents 50 μm.

Malate dehydrogenase activity in tetr apioid embryos

Because larger numbers of cytB-induced tetraploid embryos could be obtained these were used in a pilot study on the effect of tetraploidy on MDH activity (Table 2). In control 2N embryos, between the morula and blastocyst stage, MDH activity increased 16·2 %, from 4·939 × 10−10 to 5·740 × 10−10 moles NAD reduced/h/embryo, respectively. In cytB-4N embryos, the increase was 12·3 % while in cytB-4N embryos the increase was 23·7 %. In the case of cytB-4N embryos, the percent increase in MDH activity between the morula and blastocyst stage was the same whether such embryos were grown with or without zonae. In addition, the actual MDH activity of cytB-4N embryos at each stage was also approximately the same regardless of whether or not the zona pellucida was present. Therefore, in the ratios reported below, MDH activities of cytB-4N embryos were determined by pooling values of w/Z- and ZF-cytB-4N embryos.

Table 2.

Malate dehydrogenase activity in cytochalasin B-induced tetr apioid embryos

Malate dehydrogenase activity in cytochalasin B-induced tetr apioid embryos
Malate dehydrogenase activity in cytochalasin B-induced tetr apioid embryos

The MDH activity of cytB-4N morulae (90 h post-hCG) was 1·64 times that of cytB-4N morulae. When compared to w/Z-2N embryos, cytB-4N embryos had T54 times the 2N level of enzyme activity. Although the ratio of cytB-4N: cytB-4N differs from 2·0 with only borderline significance (Z = 1·544,0·2 > P > 0·1), the ratio of cytB-4N: w/Z-2N is significantly different from 2·0 (Z = 2·995, P < 0·05). The MDH activity of cytB-4N morulae was 0·94 times that of w/Z-2N morulae, not differing significantly from a hypothetical value of 1·0.

At the blastocyst stage (120 h p-hCG), the MDH activity of cytB-4N embryos was T49 times that of cytB-4N embryos. When compared to w/Z-2N blastocysts, cytB-4N blastocysts had T46 times the 2N level of enzyme activity. Both of these values differ significantly from the expected ratio of 2·0: 1·0 (Z = 4·094, P < 0·05; Z = 5·996, P < 0·01, respectively). The MDH activity of cytB-4N blastocysts was 0·98 times that of w/Z-2N blastocysts, not differing significantly from the hypothetical value of 1·0.

Tetraploid embryos obtained by blastomere fusion were then analysed to determine if PEG-4N embryos had a depression in MDH activity (relative to the expected doubling compared to 2N controls) similar to the depression observed in cytB-4N embryos (Table 3). In these experiments, between the morula and blastocyst stage, MDH activity in control 2N embryo increased 30·5 %, from 7·601 × 10−10 to 9·916 × 10−10 moles NAD reduced/h/embryo, respectively. Enzyme activities of diploid embryos grown with or without zonae were pooled because no differences in activity were detected between the two populations. In PEG-4N embryos, MDH activity increased 37·1 % between the morula and blastocyst stage. Due to a paucity of material, no PEG-2N embryos were analyzed at the morula stage, although MDH activities of such embryos were determined at the blastocyst stage.

Table 3.

Malate dehydrogenase activity in PEG /fusion-induced tetraploid embryos

Malate dehydrogenase activity in PEG /fusion-induced tetraploid embryos
Malate dehydrogenase activity in PEG /fusion-induced tetraploid embryos

The enzyme activity of PEG-4N morulae (90 h post-hCG) was 1·22 times that of control 2N embryos, which significantly differs from the expected ratio of 2·0 (Z = 12·978, P < 0·01). Thirty hours later, at the blastocyst stage, PEG-4N embryos had 1-29 times the enzyme activity of 2N embryos, and 0-95 times the enzyme activity of PEG-2N embryos. Both of these values differ significantly from the expected ratio (Z = 2·729, P < 0·1; Z = 16·454, P < 0·05, respectively).

Although it might have been expected that the doubled genome of tetraploid embryos would have been accompanied by a doubling in MDH activity relative to diploid control embryos, these data show that this correlation was not detected. This result was observed regardless of the means used to induce tetraploidy. Rather than a 100 % elevation in enzyme activity, in cytB-4N embryos the MDH activity was elevated by only 50 %, while in PEG-4N embryos the MDH activity was elevated by only about 20 %. In fact, in comparing PEG-4N with PEG-2N embryos, their enzyme activities at the blastocyst stage were essentially the same.

If the doubled genome had been accompanied by doubled enzyme activity, PEG-4N or cytB-4N embryos would be expected to have an enzyme activity equal to that of w/Z-2N embryos. In the case of cytB-4N embryos, this is what was observed. If one assumes that cytB-4N embryos did indeed have half the cell number of W/Z-2N embryos (see below), then each cytB-4N embryo blastomere had about 190 % the enzyme activity of a w/Z-2N embryo blastomere. Similarly, the MDH activity of cytB-4N embryos was only 1·5 times the activity of cytB-4N embryos. Again, if cytB-4N embryos did, indeed, have twice the cell number of cytB-4N embryos, then each blastomere of a cytB-4N embryo had about 130 % the enzyme activity of a blastomere of a cytB-4N embryo.

Cell number of PEG-4N embryos

Cell numbers of PEG-4N, w/Z-2N and ZF-2N morulae were determined 94 h post-hCG (Table 4) to see whether the data from the MDH assays resulted from cell loss in tetraploid embryos.

Table 4.

Cell numbers per morula*

Cell numbers per morula*
Cell numbers per morula*

PEG-4N morulae averaged 12·73 ±3·05 cells. w/Z-2N morulae at the same age had 17·73 ± 6·36 cells, while ZF-2N control embryos had 14·47 ± 4·02 cells, the total average for the control morulae being 16·14 ± 5·49 cells. The difference in cell number between tetraploid morulae and control morulae was not statistically significant (t = 3·01, P < 0·01). Even if the apparent 20% reduction in cell number was significant, it could not account for the full 40 % reduction from the expected doubling of MDH activity in quartet tetraploid embryos relative to whole diploid control embryos.

RNA synthesis in tetraploid embryos

Collectively, the results of the MDH assays and cell number determinations suggested that neither impaired viability nor reduced cell number could adequately account for the observed depression of MDH activity in PEG-4N embryos as compared to an expected level of MDH activity twice that of 2N embryos. One possible alternative explanation could have been that the depression in enzyme activity resulted from reduced RNA synthesis in the PEG-4N embryos due to gene dosage compensation. This hypothesis was tested by quantitating levels of nuclear RNA synthesis in a series of autoradiographic experiments.

The average number of grains overlying a nucleus as measured in each experiment is shown in Table 5. Because the number of grains per nucleus varied between experiments, for purposes of comparison, the ratio of PEG-4N grains per nucleus/2N grains per nucleus was determined for each experiment. In comparing PEG-4N morulae with w/Z-2N morulae (94 h post-hCG, both groups of embryos having undergone an equivalent number of cell divisions) the ratio varied between 0·85 and 1·07, the mean being 0·98. In the comparison between PEG-4N and ZF-2N morulae, the ratio of grains per nucleus was 1·25 and 1·66, the mean being 1·46. Autoradiographic analysis of PEG-2N embryos was not undertaken because of a paucity of such embryos, and because the MDH assays showed that PEG-2N embryos were similar to 2N embryos not exposed to PEG.

Table 5.

Tritiated uridine incorporation into RNA of morulae (94 h post-hCG)

Tritiated uridine incorporation into RNA of morulae (94 h post-hCG)
Tritiated uridine incorporation into RNA of morulae (94 h post-hCG)

Thus, in no case were nuclei of PEG-4N morulae observed to have twice the number of grains as nuclei of 2N morulae. Although the ratio of synthesis between PEG-4N and ZF-2N morulae was greater than that between PEG-4N and w/Z-2N morulae, in neither case was there evidence that the entire supernumerary genome was expressed. The results of these experiments are consistent with the idea that the MDH activity in PEG-4N embryos could, at least in part, be accounted for by partial dosage compensation of the supernumerary genome in the tetraploid nuclei.

In this study we found no statistically significant evidence for an effect of tetraploidy on the timing of preimplantation development in mouse embryos as measured by two parameters: (1) the time at which embryos developed a blastocoele and (2) the time at which embryos expressed a stage-dependent cell surface antigen. This lack of difference in timing of development was observed regardless of whether the tetraploid embryos were produced by PEG-mediated blastomere fusion, or by cytB incubation. Likewise varying cell number of either 2N- or 4N-embryos did not affect the timing of cavitation.

These results are in accord with earlier observations by Smith & McLaren (1977) who detected neither a cell number effect in 2N embryos nor an effect of cytB-induced tetraploidy on the timing of blastocoele formation. It should be pointed out that in the Smith and McLaren study, their cytB-4N embryos corresponded to our cytB-4N embryos.

MDH activity, which served as a parameter of embryonic metabolism, was monitored to determine whether the response of embryonic metabolism to tetraploidy might provide an explanation for the timing data. We found that cytB-4N embryos had only 1·5 times the MDH activity of 2N embryos, while PEG-4N embryos had only 1·2 times the MDH activity of 2N embryos. Thus, MDH activity, and presumably, overall embryonic metabolism, did not increase by a factor of two in response to a 2-fold increase in the embryonic genome.

These data on MDH activity might simply be a result of depressed embryo viability of 4N embryos, with perhaps, PEG reducing embryo viability more than cytB. However, although there is some difference in MDH activities between cytB-4N and PEG-4N embryos relative to their respective 2N controls, the actual MDH activity levels of cytB-4N and PEG-4N embryos are very similar. In addition, in both types of 2N embryos, the increase in MDH activity as 4N morulae developed into blastocysts was very similar in magnitude to the increase in MDH activity of 2N morulae as they developed into blastocysts. Thus, for both types of 4N embryos, the ratios of enzyme activity (cytB-4N : 2N and PEG-4N:2N) remained constant, not decreasing between the morula and the blastocyst stage as would be expected if the embryos were progressively deteriorating. If, then, tetraploidy itself impairs embryo viability and if PEG impairs viability more so than cytB, then these impairments must have remained constant over the 30 h period during which MDH activities were measured. A final argument against these data being a total result of depressed embryo viability are two observations, namely, (1) cytB-4N and PEG-4N embryos formed blastocysts at similar frequencies and (2) both types of 4N embryos had similar timing rates for blastocyst formation.

Another possible explanation for the apparent depressed level of MDH activity in 4N embryos is that measurements of enzyme activity could have been made at a point where the activity curve had reached a maximum. Only in the region of the curve where activity per unit substrate was uniformly increasing would it be possible to detect a doubling of activity. If measurements were made in the non-linear region of the activity curve, then, as more embryos were assayed per sample, activity per embryo would decrease. This, however, was not the case, since the numbers of PEG-4N embryos per sample varied between 3 and 5 with no change in calculated activity per embryo. The correlation of activity-to-embryos per sample is 0·419 for morulae (P > 0·1 that activity is independent of number of embryos) and −0·031 for blastocysts (P > 0-5 that activity is independent of number of embryos). Finally, our 2N levels of MDH activity are in good agreement with those previously reported by Brinster (1966), suggesting that our MDH data are cogent.

Another possible explanation for the MDH data is that 4N embryos had fewer cells than did 2N embryos. However, the difference in cell number between 4N- and 2N embryos was not statistically significant, and cannot, therefore, account for the MDH data.

If neither depressed embryo viability, nor errors in enzyme activity measurements, nor cell loss can account for the MDH data, could chromosomal loss or gene dosage compensation explain the observed, less than 2-fold increase in MDH activity in 4N embryos? Our chromosomal counts showed that putative 4N embryos were truly 4N with no evidence of partial chromosomal loss or 2N/4N mosaicism (these results and Eglitis, 1980). Thus, chromosomal loss is probably an unlikely reason for the MDH data.

To determine whether gene dosage compensation might account for the MDH data, RNA synthesis was measured autoradiographically in 2N and 4N embryos. The resulting grain counts indicated that the mean level of RNA synthesis in PEG-4N embryos was 1 -45 times that of ZF-2N embryos and 0-98 times that of w/Z-2N embryos. The difference in these two ratios is due to finding that the level of RNA synthesis in ZF-2N embryos was 35 % lower than that in w/Z-2N embryos. This finding could, then, be evidence for decreased viability of zone-free embryos in general, including, then, of the PEG-4N embryos whose levels of RNA synthesis were measured in these experiments. This would mean that the higher of these two ratios (1-45) is the more meaningful result. However, it is noteworthy that of all three embryo types (ZF-2N, w/Z-2N and PEG-4N), the standard error is greatest for ZF-2N embryos. It is possible, then, that the higher ratio results from a sampling error, particularly since the elevated ratio stems, in large part, from one experiment.

Regardless of whether the higher or the lower ratio is more accurate, it is clear that PEG-4N embryos do not synthesize twice as much RNA as do 2N embryos. Although the possibility remains that the RNA data result from decreased embryo viability, for the reasons discussed earlier, reduced embryo viability probably cannot explain all of the observed MDH and RNA data. These results are, however, compatible with the intriguing possibility that in PEG-4N morulae, compensation is made for at least part of the excess genetic material.

Although RNA synthesis in cytB-4N embryos was not quantitated, comparison of the MDH activities in cytB-4N embryos with that of cytB-4N embryos suggests that the excess genome was not as thoroughly compensated for in cytB-4N embryos as it was in cytB-4N embryos. This possibility is consistent with the idea that the control of gene activity is sensitive to cell number-related cell-to-cell interactions.

In 1977, Smith & McLaren concluded that the control of developmental timing most likely resided in the ratio of nuclear-to-cytoplasmic volume, or, in the number of nuclear divisions (DNA replications) that an embryo had undergone. The results here do not contradict either of these possibilities. In both PEG-4N and cytB-4N embryos, the ratio of nuclear-to-cytoplasmic volume remains the same as in 2N embryos of the same age. In blastomere fusion, as nuclear volume increases with tetraploidy, so does the cytoplasmic volume. Similarly, nuclear volume in cytB-4N embryos doubles, and, since, cell division is blocked during cytB treatment, cytoplasmic volume also doubles. Tetraploid embryos probably undergo the normal number of nuclear divisions since neither cell numbers nor ploidy are perturbed past the morula stage (Snow, 1973; Eglitis, 1980).

Because of the evidence for some degree of gene-dosage compensation in 4N embryos, our results do not permit a conclusion to be reached on the question of how great a role the embryonic genome actually has in developmental timing. However, these results do suggest that the nucleus is under tight control that may involve nuclear-cytoplasmic interactions sensitive to cell number.

We would like to thank Dr Irwin R. Konigsberg for some useful suggestions, and Dr E. E. Oliphant for his helpful criticisms during the preparation of the manuscript. Dr Patricia Rodier assisted with the statistical analyses, while Dr Donald A. Keefer helped with the autoradiographic study, to which Dr S. K. Lau lent his technical expertise. This report was submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy with the Department of Anatomy, University of Virginia. This work was supported by an NIH grant to L.M.W., NICHHD 1-RO1-11788. M.A.E. had additional support from an NIH predoctoral training grant.

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