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
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ABSTRACT |
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INTRODUCTION |
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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, 1984a), which can interfere with microtubule integrity and tubulin polymerization, causing chromosome segregation errors (Tuma and Sorrell, 1987
; Tuma et al., 1991
).
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, 1983; Kaufman and Bain, 1984a
). Alcohol given i.p. at specific times after ovulation can alter the quality of oocytes, increasing parthenogenesis (Dyban and Khozai, 1980
). Chronic ethanol consumption impairs the reproductive cycle and alters ovarian function (Van Thiel et al., 1978
; Eskay et al., 1981
; Cebral et al., 1998a
). Alcohol also produces an increased risk of infertility in women (Mello, 1988
; Grodstein et al., 1994
), altered oocyte quality (Cebral et al., 1998b
), reduced in vitro fertilization (Cebral et al., 1997
), and retarded and impaired preimplantation development in vitro (Cebral et al., 1999
).
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.
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MATERIALS AND METHODS |
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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 maltosedextrin (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., 1997, 1998a
).
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 2432 h post-HCG. The zygotes from the oviducts of each female were recovered by flushing with M2 medium (Quinn et al., 1982) and washed to remove the cumulus cells and the spermatozoa. They were placed into 100 µl of M16 medium (Whittingham, 1971
), 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 (7476 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 810 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 (9699 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 2 test. The Instat Program (GraphPAD software, San Diego, CA, USA) was used for calculations. A P < 0.05 was considered as significant.
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RESULTS |
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Table 5 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 5
(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 5
(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 5
(c)].
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DISCUSSION |
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We have found previously that 10% ethanol produced elevated fragmentation in the oocytecumulus complex recovered at 16 h post-HCG and maintained for 5 h in culture medium with or without spermatozoa (Cebral et al., 1998b, 1999
). 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., 1998a
), 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., 1999), 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, 1990; Henery and Kaufman, 1992
). 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, 1984b
). 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., 1995
), 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., 1999
), 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 embryomaternal 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.
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ACKNOWLEDGEMENTS |
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FOOTNOTES |
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