ALTERATIONS IN PREIMPLANTATION IN VIVO DEVELOPMENT AFTER PRECONCEPTIONAL CHRONIC MODERATE ALCOHOL CONSUMPTION IN FEMALE MICE

Elisa Cebral*, Andrea Lasserre, Valeria Rettori and Martha A. de Gimeno

Centro de Estudios Farmacológicos y Botánicos (CEFYBO-CONICET), Serrano 669, (1414) Capital Federal, Argentina

Received 14 December 1999; in revised form 24 February 2000; accepted 24 March 2000


    ABSTRACT
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
Although many studies have explored the effects of acute or chronic ethanol exposure during the postimplantation period on embryo/fetal development, few reports have described the ethanol effects on preimplantation embryo development. Little is known about the effects of ethanol consumption prior to gestation on embryo growth. Recently, we have shown that chronic moderate ethanol intake by prepubertal female mice reduces the ovulatory response and impairs in vitro fertilization and in vitro embryo preimplantation development. The purpose of the present work was to evaluate the effects of preconceptional chronic moderate ethanol ingestion on preimplantation embryo morphology and differentiation, the timing of cleavage and embryo growth in vivo, and to determine the time pattern in which alterations appear. Prepubertal female mice were treated with 10% (w/v) ethanol for 30 days prior to conception. After inducing ovulation on day 27 and 29 of the ethanol treatment, females were mated with control males and the day of presence of vaginal plug was day 1. On day 1, a decreased percentage of normal fertilized oocytes, elevated parthenogenetic oocyte activation and unfertilized eggs with abnormal metaphase II were found in ethanol-treated, compared to control females. On day 2, while any differences in the total percentage of 2-cell embryos were observed, the treated females had a significantly higher percentage of morphologically abnormal embryos, compared to control females. On day 3, the preconceptional consumption of ethanol produced significantly reduced percentages of compacted morulae and an increased percentage of uncompacted morulae. The total percentage of morulae in the treated females was lower than in controls. On day 4, ethanol-treated females showed significantly decreased percentages of hatched attached blastocysts and increased early blastocyst and morula percentages, compared to controls. Thus, preconceptional chronic moderate ethanol ingestion by prepubertal female mice produced retarded development, impaired blastocyst hatching, abnormal embryo morphology and embryo loss by fragmentation due to alterations induced in the female gamete.


    INTRODUCTION
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
Animal models of alcohol consumption have used high doses of ethanol at different stages of the organogenetic period in an attempt to induce the anatomical defects of the human fetal alcohol syndrome (FAS) (Weston et al., 1994Go; Abel, 1995Go). FAS can occur in infants born of mothers who are chronic heavy daily drinkers (Kaufman, 1997Go). However, the susceptibility of the preimplantation stages of embryo development to maternal alcohol exposure still remain relatively unexplored. In this regard, Checiu and Sandor (1982, 1986) reported reduced implantation with impaired oviductal embryo transport, and retarded and abnormal development of mouse embryos associated with acute maternal ethanol intoxication during the preimplantation period. But ethanol consumption by the mother both prior to and at the time of conception may be potentially even more harmful than ethanol consumption during pregnancy.

At present, it is unclear whether alcohol has actions at the cellular or subcellular level, or if the underlying mechanisms vary according to the organ system, the dose, the time and/or the duration of the exposure. Ethanol might act as a mutagen directly or indirectly through its first metabolite, acetaldehyde (Kaufman and Bain, 1984aGo), which can interfere with microtubule integrity and tubulin polymerization, causing chromosome segregation errors (Tuma and Sorrell, 1987Go; Tuma et al., 1991Go).

Most studies with animal models were conducted with high doses of ethanol in liquid diets that resulted in elevated ethanol-blood/tissue levels. However, models based on alcohol intake in drinking water were seldom used, because they produced low blood-alcohol levels. The administration of ethanol by mouth to female mice at periconception was capable of inducing a relatively high incidence of aneuploidy in the resultant zygotes (Kaufman, 1983Go; Kaufman and Bain, 1984aGo). Alcohol given i.p. at specific times after ovulation can alter the quality of oocytes, increasing parthenogenesis (Dyban and Khozai, 1980Go). Chronic ethanol consumption impairs the reproductive cycle and alters ovarian function (Van Thiel et al., 1978Go; Eskay et al., 1981Go; Cebral et al., 1998aGo). Alcohol also produces an increased risk of infertility in women (Mello, 1988Go; Grodstein et al., 1994Go), altered oocyte quality (Cebral et al., 1998bGo), reduced in vitro fertilization (Cebral et al., 1997Go), and retarded and impaired preimplantation development in vitro (Cebral et al., 1999Go).

There have been very few studies about the effects of pregestational ethanol intake on the morphology and growth of embryos. The purpose of this work was therefore to investigate if the preconceptional chronic consumption of moderate amounts of ethanol affects in vivo fertilization (the pronucleous formation and/or the nuclear status of the zygotes), and if previous intake of ethanol has deleterious effects on different stages of preimplantation embryo development in vivo.


    MATERIALS AND METHODS
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
Animals
Groups of three to four hybrid F1 mice (C57/B1 x CBA) from our colony were kept in plastic cages and maintained under controlled room temperature (25°C) and light cycles (14 h light: 10 h dark, lights on at 06:00). They were fed ad libitum with a commercial mouse chow diet (Diet No. 1, Nutrimentos S.A., Buenos Aires, Argentina). Daily caloric intake was estimated by the caloric value of the diet used (2900 kcal/kg). The amount of food consumed by female mice was determined by the difference between the offered chow and the remaining food plus the amount spilled.

Adult 75-day-old male mice (average body weight ± SEM: 29.4 ± 0.68 g) were used. Females were immature (prepubertal, 30-day-old, average body weight: 17.6 ± 0.35 g) at the start of the ethanol administration period. Control female mice were the same age and weight as the ethanol-treated mice at the beginning of the ethanol treatment.

Ethanol treatment
Immature female mice were treated with 10% (w/v) ethanol in drinking water for 30 days. The ethanol exposure was stopped at the beginning of pregnancy. Controls received water with maltose–dextrin (3.8 kcal/g) in a proportion of 56 g to 300 ml water, to be isocaloric with 10% ethanol. The body weights were recorded daily throughout the treatment in all females used. The amount of daily liquid intake was determined by volume differences between the offered and the remaining volume. Calories derived from ethanol were estimated as 7.1 kcal/g. From these data, daily patterns of caloric intake and the percentage of ethanol-derived calories (% EDC) were determined in five female mice for each experimental group.

The effects of preconceptional chronic moderate ethanol ingestion were examined on in vivo fertilization (day 1, pronucleus formation) and on in vivo development (from day 2 to day 4).

Blood-ethanol measurement
A group of five immature female mice chronically treated with ethanol as described above were decapitated and trunk blood was collected into heparinized Eppendorf tubes at 06:00 on day 30 of treatment. Samples were maintained at 4°C, for blood-ethanol determinations within 4 h of collection. Ethanol was measured by gas chromatography as described previously (Cebral et al., 1997Go, 1998aGo).

In vivo fertilization
Female mice were induced to superovulate with 10 IU of pregnant mare's serum gonadotrophin (PMSG, Sigma Chemical Co., St Louis, MO, USA) given at 13:00 on day 28 of the ethanol treatment and 10 IU of human chorionic gonadotrophin (HCG, Sigma) 48 h later (day 30). One female was than caged immediately with one adult non-treated male, previously isolated. On the morning of the next day, mating was confirmed by the presence of the vaginal plug (day 1). The following groups were studied (1) control group: control females mated with control males; and (2) ethanol-treated group: ethanol-treated females mated with control males.

In vivo fertilization was evaluated at 24–32 h post-HCG. The zygotes from the oviducts of each female were recovered by flushing with M2 medium (Quinn et al., 1982Go) and washed to remove the cumulus cells and the spermatozoa. They were placed into 100 µl of M16 medium (Whittingham, 1971Go), supplemented by 3% bovine serum albumin (BSA) and overlaid with mineral oil. The medium contained the vital fluorocrome Hoechst 33342 (0.5 µg/ml, Sigma). The embryos and/or oocytes were incubated for 1 h, washed and mounted for observation under a fluorescence microscope. They were classified as fertilized oocytes when they presented the second polar body (II PB) with two pronuclei (2PN) (fertilized normally). The triploid oocytes (polyspermic, with II PB and 3 PN) and oocytes with II PB and 2 PN were also considered as fertilized. Unfertilized activated oocytes were parthenogenetically activated and appeared with: II PB and 1 PN, II PB and dispersed chromosomes (DC) in the cytoplasm (metaphase plate disruption), 0 PB (without second polar body) and 1 PN and 2-cell embryo from immediate cleavage. Unfertilized intact oocytes were oocytes with normal metaphase II plate (Me II) or oocytes with abnormal metaphase (disruption of metaphase plate with decondensed and DC). Fragmented and/or necrotic (lysed) oocytes were recorded as degenerated.

In vivo development
Females were induced to superovulate with PMSG and HCG (10 IU) and caged with one male previously isolated at the time of HCG injection, as described before. On the following morning, mating was confirmed by the presence of the vaginal plug (day 1 of pregnancy). The same groups as indicated previously were studied.

(a) Collection of embryos. Females were killed by cervical dislocation at varying days of in vivo development. The eggs, embryos and other elements (intact, fragmented and/or dead oocytes, empty zona pellucida) from each female were recovered from the oviducts and/or uteri by flushing with M2 medium. The cells were washed to remove pieces of tissue and placed in a 100-µl drop of M16 medium supplemented with BSA 3% and covered with mineral oil, for examination under an inverted phase contrast microscope and for further studies. Results were expressed as the mean percentage of embryos calculated with the total number of elements (intact or degenerated eggs, embryos and/or oocytes) recovered per female.

(b) Evaluation of general development. On day 2 (48 h post-HCG), 2-cell embryos were quantified from the oviducts. Other elements were recorded: II PB (one cell with second polar body), 4-cell embryos (embryos derived from activated oocytes or immediatelly cleaved embryos), oocytes without zona pellucida and/or empty zona pellucida.

On day 3 (74–76 h post-HCG), embryos recovered from oviducts were at the morula stage (8-cell uncompacted and compacted embryos). Other elements can also appear: 1-, 2- or 4-cell embryos and/or unfertilized eggs.

On day 4 (96 and 99 h post-HCG), the following were obtained: morulae, no hatched blastocysts (non-extruded embryos from the zona pellucida) and hatched blastocyst (zona-free embryos). The blastocyst differentiation grade was assessed by quantification under phase-contrast microscopy as the very early blastocyst (embryo with small vesicles in blastomeres), early blastocyst (with an initial visible blastocele), expanded blastocysts (with expansion of the cavity), hatched early blastocysts (zona-extruded blastocysts with visible cavity), and hatched attached blastocysts (extruded from the zona pellucida without a visible cavity or implanting blastocyst appeared). This embryo classification was completed and confirmed by counting the number of nuclei/embryo, as indicated above. Non-fragmented embryos recovered from oviducts or uteri were quantified to evaluate the utero-tubaral distribution of embryos 99 h post-HCG.

(c) Evaluation of embryo quality. Embryo quality was determined by the following parameters: extracellular fragmentation (presence of small clumps of cytoplasm surrounded by membrane in the perivitelline space); no intact blastomeres (with necrotic signs or lysis); blastomeres with unequal size; decompacted blastomeres in compacted morulae; abnormal cavitation (two or more blastoceles); vesicles in ectoderm; small inner cell mass. When the embryo presented with more than 50% of fragmentation, it was considered to be fragmented. Embryos with high rates of lysis were considered to be dead embryos. Both fragmented and dead embryos were classified as ‘degenerated’.

(d) Evaluation of embryo growth. On days 3 and 4, the number of cells/embryo was determined by counting the nuclei of individual embryos recovered from each group, according to the air-drying technique of Tarkowski (1966). Briefly, the embryos were swollen in drops of 0.9% sodium citrate for 8–10 min and transferred to clean slides to fix for 30 s by dropping a single drop of methanol : acetic acid (3:1 v/v) over the embryos, and air dried. The cytoplasm of the embryos was dispersed by this technique leaving only chromatin behind. The cells were stained with Giemsa solution (2:1, v/v) for 10 min and the nuclei were examined under light microscopy.

On day 3, the mean number of cells/embryo, mean number of mitoses/embryo and the percentage of embryos with mitotic cells were quantified in the embryos recovered in each group. The mitotic index (MI) was calculated as the total number of metaphases/total number of cells. On day 4 (96–99 h post-HCG), the distribution of nuclei/embryo was assessed in a total of 56 embryos from seven control females and 54 embryos from seven ethanol-treated females. The result was expressed as the percentage of embryos vs the number of cells/embryo range.

Statistics
The mean percentages of developed embryos and SEM were calculated with the total elements recovered/treated female (intact embryos, eggs, zygotes, oocytes and/or others) (n = no. of females used). The data from these two groups were analysed by Student's t-test. The percentages of embryos in the evaluation of the embryo growth were examined by the {chi}2 test. The Instat Program (GraphPAD software, San Diego, CA, USA) was used for calculations. A P < 0.05 was considered as significant.


    RESULTS
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
Average daily intakes of food, fluid, and ethanol in females, expressed in g, ml, and/or kcal/mouse respectively, are shown in Table 1Go. These data were recorded in five female mice for each group. The percentage of ethanol derived calories (% EDC) was 26.8%. The body weight reached at the end of the treatment was similar in the control and ethanol-treated females used in all studies. The total caloric intake was similar in both groups. Blood-ethanol levels on the morning of the last day of treatment were 48.1 ± 1.5 mg/dl.


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Table 1. Daily intake of food, fluid, and percentage of ethanol-derived calories in female mice chronically treated with 10% ethanol
 
In vivo fertilization
Table 2Go shows that chronic 10% ethanol treatment produced a significantly decreased percentage of normal fertilized oocytes (with II PB and 2 PN), as compared to the control females [P < 0.01, Table 2Go(a)]. Unfertilized activated eggs (with II PB and 1 PN) were present significantly more in the treated females than in the controls [P < 0.01, Table 2Go(b)], but also a significantly increased percentage of oocytes with 0 PB and 1 PN (diploid eggs) could be seen in the treated group [P < 0.05, Table 2Go(b)]. A significantly elevated percentage of unfertilized intact oocytes with abnormal metaphase plates was found in the treated females, as compared to the controls [P < 0.05, Table 2Go(c)]. The total percentage of fragmented and/or dead oocytes was similar between the control and the ethanol-treated group (data not shown).


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Table 2. Effect of chronic 10% ethanol intake on in vivo fertilization
 
In vivo development
On day 2, there was a similar percentage of 2-cell embryos in both control and ethanol-treated females [Table 3Go(a)], but the percentage of morphologically abnormal 2-cell embryos was significantly increased in the treated, compared to the control, group (P < 0.05). No significant group differences were observed in other cells [Table 3Go(b)].


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Table 3. Effects of 10% ethanol consumption on day 2
 
On day 3 (76 h post-HCG), the total percentage of morulae was significantly decreased in the ethanol-treated, as compared to the control, females [P < 0.01, Table 4Go(a)]. The compacted morula stage was significantly reduced (P < 0.001) and uncompacted morulae were increased (P < 0.05) in the ethanol-treated group compared to the control group. The percentage of degenerated embryos and others (1-, 2- or 4-cell embryos, unfertilized eggs) recovered from the oviducts was significantly increased in the ethanol-treated females compared to the control females [P < 0.05 and P < 0.01, Table 4Go(b)]. The growth of embryos was analysed and the mitotic cells were assessed to evaluate the timing of division. The mean number of cells/embryo was significantly decreased in the ethanol-treated, as compared to the control, group [P < 0.05, Table 4Go(c)]. The treated females presented a significantly higher mitotic index than the control females (P < 0.05) [Table 4Go(c)].


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Table 4. Effects of 10% ethanol consumption on day 3
 
On day 4, the effects of 10% ethanol consumption on the in vivo development was first studied at 96 h post-HCG. In the control females, almost all embryos were at the hatched blastocyst stage and very few embryos were still morulae, whereas in the ethanol-treated females, the percentage of hatched blastocysts was significantly decreased (P < 0.05) and the percentage of morulae increased (P < 0.05) (data not shown). In the ethanol-treated females, the mean total percentage of embryos/female was significantly reduced compared to controls (59.8 ± 11.2 vs 94.6 ± 3.6, P < 0.01) (data not shown).

Table 5Go shows the embryo stages developed on day 4 at 99 h post-HCG. In the ethanol-treated group, the percentage of morulae and un-hatched blastocysts was significantly increased compared to the control group [P <0.05, Table 5Go(a)], but the hatched blastocyst and the total number of blastocysts was significantly decreased in the ethanol-treated group, compared to the control group (P <0.05). When the blastocyst differentiation was analysed, the percentage of early blastocyst was increased significantly (P <0.01), and the percentage of attached blastocyst stage was significantly reduced in the ethanol-treated females, compared to the control females [P < 0.01, Table 5Go(b)]. The total percentage of fragmented and/or dead embryos and the percentage of abnormal embryos were significantly higher in the ethanol-treated, than in the control, group [P < 0.01 and P < 0.05, Table 5Go(c)].


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Table 5. Effects of 10% ethanol consumption on day 4 of development: embryos from 99h post-HCG
 
The utero-tubaral distribution of embryos recovered at 99 h post-HCG is shown in Table 6Go. As can be seen, in the ethanol-treated females, almost all non-fragmented embryos of the total percentage of embryos recovered (71.5%) were found in the uteri, and the percentage was significantly reduced, compared to the control females (P < 0.05). The rest of the embryos were found in the oviduct tract, but we did not observe significant differences between the control and the ethanol-treated groups.


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Table 6. Utero-tubaral distribution of embryos
 
Embryo growth was analysed by counting the number of cells/embryo recovered on day 4. This evaluation allowed the correct classification of embryos in stages of morulae (less than 32 cell embryos), very early blastocyst, early blastocyst, expanded blastocyst, and hatched blastocyst (both early and attached) (Table 7Go). In the ethanol-treated females, the percentage of embryos with 31–40 cells/embryo (very early and early blastocyst) was significantly increased, compared to the controls (P < 0.05) (Fig. 1Go). The percentage of embryos with more than 60 cells (preferentially hatched blastocysts) was significantly lower in the treated group than in the control group (P < 0.001) (Fig. 1Go).


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Table 7. Stages of embryo development in vivo in relation to the number of nuclei on day 4
 


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Fig. 1. Embryo growth: number of cells/embryo on day 4.

The total number of nuclei/embryo was determined by the Tarkowski (1966) method. Columns represent the percentage of embryos in each range of number of cells assessed. Black columns: control females, white columns: ethanol-treated females mated with control males. *P < 0.05,

***P < 0.001, {chi}2-test.

 

    DISCUSSION
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
We postulated that prolonged ingestion of moderate doses of ethanol by young female mice prior to gestation could affect in vivo fertilization and the preimplantation in vivo development. We considered it relevant to evaluate the zygotes after mating, on day 1 of development, in order to study not only the in vivo fertilization rates, but also the nuclear status of the eggs. We found that the normal fertilized oocyte rates were reduced in the treated females, because elevated parthenogenetic activation values were observed. These results are in agreement with our previous studies, in which chronic consumption of 10% ethanol produced a high percentage of oviductal activated oocytes (1 PN) in the cumulus–oocyte complexes at 16 and 20 h post-HCG without fertilization (Cebral et al., 1998bGo). Moreover, we have seen that chronic moderate ethanol exposure produced a reduced percentage of fertilized oocytes after in vitro insemination, because there was an elevated percentage of haploid oocytes (1 PN) (Cebral et al., 1999Go). It has been suggested that ethanol can induce parthenogenetic activation of oocytes (meiosis resumption) (Kaufman, 1978Go) by releasing internal calcium pools (Whittingham, 1980Go). Calcium signalling during parthenogenetic egg activation was postulated to affect preimplantation development, even several cell divisions after Ca2+ oscillations occur (Vitullo and Ozil, 1992Go; Bos-Mikich et al., 1997Go). Further, calmoduline activation by Ca2+ initiates several cellular events (Lu and Means, 1993Go), including altered gene expression (Hama et al., 1995Go). Prolonged ethanol exposure could not only trigger Ca2+ signals and activation of the oocyte, but also alter gene transcription, as occurs in neuronal (Wilke et al., 1994Go; Ghosh and Greenberg, 1995Go) and non-neuronal (Fatayerji et al., 1996Go) cell types. Ethanol treatment for a long period of time was capable of inducing abnormal metaphase plates and therefore chromosomes could be dispersed in the cytoplasm of intact oocytes after fertilization. Some mechanism(s) of these ethanol effects may be related to the involvement of abnormal chromosome segregation (Kaufman, 1983Go). Multipolar spindles and some microtubules at the periphery of the spindle, that may be associated with disorganized alignment of the chromosomes, were encountered following ethanol exposure (O'Neill and Kaufman, 1989Go). High incidence of aneuploidy due to ethanol intake around the ovulation time (Kaufman and Bain, 1984aGo; Kaufman, 1985Go), interference with cell division (Kaufman, 1997Go), and cytoskeletal structure and function of the spindle apparatus (Kaufman, 1985Go; O'Neill and Kaufman, 1989Go), disrupted regulation of normal calcium concentrations in the proximity of the spindle apparatus during cytokinesis (Kaufman, 1997Go) is also attributed to ethanol exposure.

We have found previously that 10% ethanol produced elevated fragmentation in the oocyte–cumulus complex recovered at 16 h post-HCG and maintained for 5 h in culture medium with or without spermatozoa (Cebral et al., 1998bGo, 1999Go). In the present study, the treated females did not show increased fragmentation rates at 32 h post-HCG. Perhaps, even when blood-ethanol levels of 50 mg/dl were considered by us to be toxic to the ovarian oocyte (Cebral et al., 1998aGo), the oviductal environment could protect against increased fragmentation in the eggs. This is in agreement with our previous study which showed that the percentage of fragmented oocytes recovered from the ampullae 20 h post-HCG was similar in both groups. We believe that the altered oocyte quality and fertilization are more clearly manifested in in vitro conditions. Moreover, the oocyte susceptibility to the deleterious effects of ethanol for increasing apoptosis is more evident by in vitro manipulation of the gamete.

We investigated whether ethanol treatment could alter the embryo growth and/or morphology at earlier days of development in vivo. On day 2, although development of 2 cell embryos was similar in both groups, embryos from ethanol-treated females presented different blastomere size or absence of polar body or fragments in the perivitelline space. These abnormal embryos could undergo arrest later, because on day 3, 2- or 4-cell embryos were also observed. However, since the percentage of 2-cell embryos developed in vivo was not found to be reduced, compared to controls, but significantly reduced when developed in vitro (Cebral et al., 1999Go), we believe that the embryos from alcoholic females have severely impaired development in vitro.

On day 3 of gestation, a delayed development accompanied by augmented embryo fragmentation was also seen, since reduced numbers of compacted morulae and increased numbers of uncompacted embryos were found in the ethanol-treated females. The decreased number of cells/embryo and the elevated mitotic index suggest that even the smaller morulae from ethanol-treated females had cells undergoing division and therefore late cell division. It has been observed that haploid or diploid parthenogenetic embryos can develop more slowly than control fertilized diploid ones (Kaufman, 1990Go; Henery and Kaufman, 1992Go). We think that this delay is due to the nuclear status of embryos induced earlier by ethanol exposure. This is also suggested by the observation of the nuclear status of the oocytes post in vivo fertilization (day 1). Moreover, previous reports have demonstrated that monosomic or trisomic embryos induced by maternal exposure to ethanol are capable of reaching at least the morula stage (Kaufman and Bain, 1984bGo). Indeed, when embryo growth was explored 96 and 99 h post-HCG (day 4), we found that the ethanol-treated females had reduced quantities of embryos (particularly the blastocyst stages), despite the similar rate of recuperation, compared to control females. A high number of these embryos showed morphological abnormalities, such as smaller size, presence of vesicles in the cytoplasm or fragments in several blastomeres and/or in the perivitelline space. We hypothesized that these embryo abnormalities may be caused by the prolonged pregestational ethanol consumption. Moreover, another adverse effect of ethanol could be the induction of apoptotic processes later in the development via the increase of cell fragmentation, as seen in the treated females. We think that morphologically abnormal embryos died later, because of fragmentation or cytoplasmic lysis, and that only the completely normal embryos survived and were implanted. It was suggested that parthenogenetic embryos can undergo apoptosis (Takase et al., 1995Go), although little is known about the mechanisms underlying the degeneration of oocytes and embryos. Since only a few embryos reached the hatched blastocyst stage, we believe that ethanol consumption could impair not only the hatching process but also the development of competent implanting blastocysts. Furthermore, the lower number of cells in the embryos from the ethanol-treated females confirms reduced blastocyst development. These results suggest that severe embryo growth retardation could be induced by pregestational ethanol ingestion, and that these effects can be detected later in development, even when ethanol consumption is stopped. In consequence, the implantation might be reduced or delayed in ethanol-treated females. We have found similar results in the in vitro development model (Cebral et al., 1999Go), when not only the process of blastocyst expansion was impaired or retarded, but also the total extrusion of the embryo from the zona pellucida was strongly affected by ethanol consumption.

The total percentage of non-fragmented/dead embryos recovered from each female was analysed for utero-tubaral distribution on day 4. The observation of reduced quantities of embryos in the uteri and a few in the oviduct tract in ethanol-treated females suggests retarded embryo transport and therefore impaired functioning of the maternal tissues. We believe that preconceptional ethanol consumption adversely affected the embryo and its capacity of development in vivo and in vitro, the functioning of maternal genital tract, and the embryo–maternal communication necessary for implantation.

In conclusion, this study shows that several stages of embryos developed in vivo were affected by chronic moderate ethanol ingestion when female mice were prepubertal and were treated before gestation. The retarded embryo development, morphological abnormalities, embryo losses, and impaired fertilization could be consequences of abnormal pronuclei formation due to impaired chromosome segregation during oocyte maturation.


    ACKNOWLEDGEMENTS
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
The authors thank Dr Marcelo Viggiano for revising this paper and for suggestions regarding the statistical analysis. This work was supported by Grant PIP 4076 from the Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET) and Bago S.A. (Argentina).


    FOOTNOTES
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
1 *Author to whom correspondence should be addressed. Back


    REFERENCES
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
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