ToxSci Advance Access originally published online on May 26, 2006
Toxicological Sciences 2006 92(2):416-422; doi:10.1093/toxsci/kfl024
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Ethanol-Induced Fetal Dysmorphogenesis in the Mouse Is Diminished by High Antioxidative Capacity of the Mother
Department of Medical Cell Biology, Biomedical Center, Uppsala Universitet, SE-751 23 Uppsala, Sweden
1 To whom correspondence should be addressed at Department of Medical Cell Biology, Biomedical Center, Uppsala Universitet, PO Box 571, SE-751 23 Uppsala, Sweden. Fax: +46-18-550-720. E-mail: parri.wentzel{at}medcellbiol.uu.se.
Received March 25, 2006; accepted May 16, 2006
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Intrauterine exposure to ethanol causes embryonic and fetal maldevelopment. Oxidative stress in mother and offspring has been suggested to be part of the teratogenic mechanism of ethanol. Here we aimed to assess the importance of maternal and fetal antioxidative capability for the risk of dysmorphogenesis in the offspring. We used male and female mice with different levels of superoxide dismutase (SOD) activity—wild-type (WT) mice, mice with a targeted SOD mutation (KO, decreased CuZnSOD mRNA), and mice transgenic for SOD (TG, increased CuZnSOD mRNA). Female WT, KO (heterozygous), and TG (heterozygous) mice were given drinking water containing 20% ethanol before and throughout gestation. Non–ethanol-exposed WT, KO, and TG mice served as controls. The female mice were mated with males with identical genotype, and the pregnancy was interrupted on gestational day 18 when the offspring was evaluated and genotyped. Fetal hepatic isoprostane (8-epi-PGF2
) levels were measured to assess the degree of fetal oxidative stress. Exposure to 20% ethanol decreased fetal weight by 9–13% in the three groups. Ethanol exposure roughly doubled the rates of maldeveloped WT and KO offspring but did not affect TG offspring. The fetal hepatic levels of 8-epi-PGF2 were increased in the ethanol-exposed WT and KO mice but not in ethanol-exposed TG mice. Ethanol exposure preferentially damaged WT fetuses in pregnant KO mice, whereas no such effect was found in the litters of ethanol-consuming TG mice. Administration of ethanol to pregnant mice disturbs embryogenesis by oxidative stress, and the adverse effects are more pronounced in offspring of mice with low antioxidative capacity.
Key Words: ethanol; mouse; transgene; targeted mutation; oxygen radical; isoprostane; 8-epi-PGF2.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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It is well established that ethanol consumption by the mother disrupts fetal development in the human (Jones et al., 1973
; Lemoine et al., 1968) and in several species of experimental animals (Goodlett and Horn, 2001; Wang et al., 2006). The precise reasons for the disturbed embryonic development have not yet been clarified.
Several suggestions have gained experimental support, indicating a putative multifactorial etiology. Thus, previous studies have supported disturbed glucose, protein and DNA metabolism (Miller and Dow-Edwards, 1988; Snyder et al., 1986), impaired neurogenesis (migration and neurite outgrowth) (Lindsley et al., 2003; Ma et al., 2003; Zhou et al., 2001), and inhibition of L1 CAM (Armant and Saunders, 1996; Charness et al., 1994) as ethanol-induced teratological mechanisms. Furthermore, decreased expression of the sonic hedgehog gene (Ahlgren et al., 2002), altered expression of genes involved in facial formation (Da Lee et al., 2004), as well as outright cell damage (increased apoptosis) (Cartwright et al., 1998; Ramachandran et al., 2003) have been suggested by previous experimental studies.
Enhanced production of reactive oxygen species (ROS) has also been suggested to be involved in the teratogenic process of ethanol-exposed pregnancy in experimental animals (Chen and Sulik, 1996; Davis et al., 1990; Devi et al., 1996; Guerri et al., 1994; Henderson et al., 1995, 1999) and pregnant women (Kay et al., 2000). The mechanisms by which ethanol-induced ROS production disrupts fetal cells may be related to membrane damage (altered fluidity and interrupted transport systems) (Henderson et al., 1999; Polache et al., 1996). In support of the notion of ROS-mediated developmental disturbances is treatment with antioxidative compounds which have been successful in preventing ethanol-induced damage to embryos both in vitro (Kotch et al., 1995; Peng et al., 2005) and in vivo (Cano et al., 2001; Chen et al., 2004; Satiroglu-Tufan and Tufan, 2004). Moreover, embryonic and fetal cells exposed to ethanol in vitro can be saved by supplementation of antioxidants (Henderson et al., 1999; Mitchell et al., 1999). Furthermore, Xenopus embryos with genetically enhanced oxidative defense (transgenic for catalase and peroxiredoxin) tolerate ethanol in vitro without developing (eye) malformations (Peng et al., 2004). In fact, the available experimental evidence in favor of antioxidative treatment has led to suggestions of prophylactic supplementation of ethanol-consuming pregnant women with antioxidative compounds, such as large doses of vitamin E and vitamin C (Cohen-Kerem and Koren, 2003).
Against this background, our aim in the present study was to investigate whether developmental damage in offspring of mice subjected to chronic ethanol consumption may be modulated by varied maternal and fetal antioxidative capacity. To monitor the presence and degree of oxidative stress in fetal tissues, we estimated the concentration of the isoprostane 8-epi-PGF2 (Morrow et al., 1990) in the liver of the offspring.
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MATERIALS AND METHODS |
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Animals.
We used three types of mice for the study (all obtained from Jackson Laboratory, Bar Harbor, ME): C57/Bl/6J mice (denoted wild type, WT), B6;129S7-Sod1tm1Leb/J with a targeted mutation of CuZn superoxide dismutase (SOD) (Matzuk et al., 1998
) (denoted KO); and C57BL/6-Tg(SOD1)3Cje/J mice transgenic for hCuZnSOD (Epstein et al., 1987) (denoted TG). The homozygous KO mice do not express a functional CuZnSOD protein at all. The total SOD activity in erythrocytes decreases by 40% in heterozygote KO mice and by 65% in homozygote KO mice. Likewise, the total SOD activity increases by 60% in the heterozygote TG mice and by 100% in the homozygote TG mice (SOD Assay Kit #5016, Cell Technology, Mountain View, CA). The mice were housed at the Laboratory Animal Resources of the Biomedical Center in Uppsala and subjected to a 12-h dark /12-h light cycle. All mice had free access to drinking water (with or without ethanol) and pelleted food R36 (Lactamin AB, Stockholm, Sweden).
Dietary treatment.
Before pregnancy, female mice were given drinking water with successively increased concentration of ethanol during 2 weeks, starting at 5% and ending at a concentration of 20% (vol/vol). The drinking water was also supplemented with 50 g/l glucose (to mask the ethanol taste) in all bottles, irrespective of whether or not they contained ethanol.
Pregnancy.
The female mice serving as controls (denoted CWT, CKO, and CTG) as well as the mice drinking ethanol-containing water (denoted EWT, EKO, and ETG) were mated overnight with normal male mice of the same strain and genotype, i.e., WT male x WT female, heterozygote KO male x heterozygote KO female, or heterozygote TG male x heterozygote TG female. Day 0 of pregnancy was defined as the day sperms were found in a vaginal smear. On gestational day 18 the pregnant mouse was weighed, and a blood sample was taken from the tip of the tail to determine concentrations of ethanol (mixed with heparin and stored at – 20°C before analysis, see below) and glucose (measured immediately with a MediSense glucose sensor, Abbott Scandinavia AB, Solna, Sweden). The pregnant mouse was killed by cervical dislocation; the litter was dissected out, weighed, and examined for the occurrence of fetal malformations and resorptions. Placental weight and litter position of each fetus were also recorded (the two positions at both ends of each uterine horn were denoted "periphery," and the status of these fetuses was compared to that of all intermediate positions, denoted "central," see below), and the tail tip was taken from each fetus to enable genotyping. If the offspring was resorbed, a piece of fetal tissue was collected and used for genotyping. From each litter, one or two fetal livers were dissected, immediately frozen, and stored at – 80°C until analyzed for the isoprostane 8-epi-PGF2 and protein content.
Measurement of ethanol.
Ultrapure water was used in all buffers and dilutions of standards and samples, e.g., filtered deionized water (Milli-Q plus, Millipore AB, Stockholm, Sweden). The blood was thawed and analyzed in accordance with the method by Cornell and Veech (1983). Briefly, 100 µl blood was mixed with 800 µl ice-cold perchloric acid (0.33M), the sample was centrifuged (3000 rpm) for 5 min, and the deproteinized supernatant was saved for further analysis. Ethanol standard samples were prepared by mixing absolute ethanol and ultrapure water to yield a 0.4% (87.0mM) ethanol stock solution. This solution was serially diluted to yield a set of seven standard samples in the range of 1.4–87.0mM ethanol. Absorbance was measured at 340 nm wavelength (UV mini 1240, Shimadzu, Stockholm, Sweden) on 3 µl standard or test sample, to which were added 100 µl Tris-lysine buffer, 91 µl ultrapure water, and and 6 µl ß-NAD (Sigma-Aldrich Sweden AB, Stockholm, Sweden). After an initial absorbance reading, 2 µl alcohol dehydrogenase (ADH, EC 1.1.1.1
[EC]
, Sigma-Aldrich Sweden AB) from a stock solution of 9000 kU/l ADH in PBS was added to each sample, and absorbance was remeasured after 20–30 min. The difference in absorbance, which is proportional to the NAD+ consumed by the reaction, was used in the calculations as an estimate of ethanol concentration:
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Genotyping offspring of KO and TG mice.
To establish the genetic identity of the offspring of the KO and TG mice, a piece of the tail of fetuses or tissue samples of resorbed offspring was obtained and placed in an Eppendorf tube containing Tris buffer (50mM TrisBase, 100mM EDTA, 100mM NaCl, 20% SDS, pH 7.5) with proteinase K and incubated at 55°C in a water bath overnight. Next morning the tubes were vortexed thoroughly and centrifuged for 10 min. We transferred 300 µl of the supernatant to a new tube containing 200 µl isopropanol. Next, the tubes were gently shaken and centrifuged (13,000 x g, 10 min). If the DNA did not precipitate as a white shadow, the isopropanol was removed and replaced by 1 ml of 70% ethanol. The tubes were subsequently centrifuged (13,000 x g, 5 min), and the supernatant (ethanol or isopropanol) was discarded. The pellet was dried and resuspended in 50 µl RNase-free water. Subsequently, the isolated DNA was subjected to RT-PCR (with the primers for the PgkHPRT minigene: TgTTCTCCTCTTCCTCATCTCC, forward; ACCCTTTCCAAATCCTCAgC, reverse, and exon 2 of CuZnSOD: TgAACCAgTTgTgTTgTCAgg, forward; TCCATCACTGGTCACTAGCC, reverse) to genotype KO offspring (with the primers: WT: CTAggCCACAgAATTgAAAgATCT, forward; gTAggTggA AATTCTAgCATCATCC, reverse; TG: CATCAgCCCTAATCCATCTgA, forward; CgCgACTAACAATCAAAgTgA, reverse) to genotype TG offspring.
Measurement of the isoprostane 8-epi-PGF2.
Estimations of 8-epi-PGF2 were performed with the aid of an enzyme immunoassay in accordance with the instructions from the manufacturer (Cayman Chemical Co, Ann Arbor, MI). Briefly, livers from mouse offspring at gestational day 18 were homogenized on ice in 1 ml 0.1M phosphate buffer, pH 7.4, containing 1mM EDTA and 10µM indomethacin. Equal volume of 15% (wt/vol) KOH was added to each liver homogenate, and the samples were incubated at 40°C for 60 min. Next, two to four volumes of ethanol were added to each sample, which was vortexed, incubated at 4°C for 5 min, and centrifuged for 10 min (1500 x g). The supernatant was decanted into a clean tube, the ethanol was evaporated, and the samples were acidified (pH 4) by the addition of 30% acetic acid. Samples were then passed through a C-18 SPE-Cartridge (Supelco DSC-18, Bellefonte, PA), which had been activated previously by rinsing with methanol followed by ultrapure water (Milli-Q plus, Millipore AB). After passing the sample, the cartridge was rinsed with ultrapure water followed by HPLC-grade hexane (Sigma-Aldrich Sweden AB). The isoprostane was then eluted with 5 ml ethyl acetate containing 1% methanol (Sigma-Aldrich Sweden AB), which was subsequently evaporated in a vacuum centrifuge (Speed Vac, SVC 100). The samples were dissolved and analyzed in a spectrophotometric plate reader (IEMS Lab System, Helsinki, Finland) at a wavelength of 405 nm. The protein content of the liver samples was estimated by the method of Lowry et al. (1951) using bovine serum albumin as a standard.
Ethical and statistical evaluation.
All animal procedures were performed according to the "Guide for the Care and Use of Laboratory Animals" (NIH, 1985) and approved by the Animal Ethics Committee of the Medical Faculty of Uppsala University. Comparisons between different experimental groups were based on individual pregnant mice or litter means, with the exception of fetal dysmorphogenesis (malformations and resorptions), where the data from all pregnancies in a particular experimental group were pooled. Differences between means were evaluated by Student's two-tailed t-test or ANOVA, where the applied post hoc test was Fisher's Protected Least Significant Difference at the 95% significance level. The rates of fetal dysmorphogenesis were compared with chi-square statistics. A value of p < 0.05 was considered to denote a significant difference between groups. The calculations were performed with the aid of the Macintosh version of the statistical program Statview.
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RESULTS |
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Random sampling of the drinking bottles yielded a mean consumption of 4–5 ml water and 3 g pelleted chow per mouse and day, with no apparent differences between the groups, thus yielding a daily mean consumption of about 25 g ethanol/kg body weight during gestation.
On gestational day 18, the maternal blood glucose levels did not differ between the groups. In the three ethanol-consuming groups, the maternal serum levels of ethanol were in the range of 4.5–5.6 mmol/l and did not differ between the ethanol-exposed groups, all of which were increased compared to non–ethanol-consuming mice (Table 1). Ethanol consumption decreased maternal weight in the EKO group compared with the weight of the pregnant CKO mice (Table 1). Fetal weight was decreased in all ethanol-consuming groups, whereas placental weight was increased only in the EWT group (Fig. 1).
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n The fetal outcome was severely affected by ethanol exposure since the proportions of malformed and resorbed fetuses were increased in the EWT and EKO mice by 90% and 119%, respectively; however, this was not the case in the ETG mice (17% difference, Fig. 2). The hepatic isoprostane concentration was increased in the offspring of the EWT and EKO mice but not in the ETG group (Fig. 3).
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Analysis of the fetal intrauterine position revealed that both ends of the uterine horn (periphery) displayed increased occurrence of resorptions and malformations in the ethanol-exposed KO group but not in the EWT and ETG mice. In the latter group, the changes tended to be reversed (Fig. 4).
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Ethanol exposure did not affect the distribution of genotypes in the different groups (Fig. 5). In contrast, ethanol caused a marked change in the distribution of genotypes among maldeveloped offspring in the EKO group, where the WT offspring was markedly overrepresented (Fig. 6). In the transgenic group, there was no effect of ethanol exposure on the distribution of genotypes in the maldeveloped offspring. Furthermore, when the body weight of the WT offspring (which should be genetically similar in all litters, although not completely identical) was compared between all the groups, it was apparent that maternal ethanol consumption decreased fetal body weight by 10% and 11% in the WT and TG litters, respectively (Fig. 7), whereas the WT placental weight was increased in the EKO and ETG litters (by 20% and 30%, respectively, Fig. 8).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The major finding of this study was that enhanced antioxidative capacity in the pregnant animal, manifested by increased expression of the CuZnSOD gene, can partly prevent ethanol-induced dysmorphogenesis in the offspring. The genotype of the fetus, on the other hand, seems to be less important for the outcome of ethanol-exposed pregnancy.
The EKO animals displayed the most hostile environment, and the ETG animals the most supportive environment, as evidenced by the increased and unchanged rates of dysmorphogenesis, respectively. It is tempting to assume that the difference in maternal (anti)oxidative status may be of primary importance for the differences in outcome. However, other genetic and metabolic effects caused by the different doses of the CuZnSOD genes could also be of developmental importance, both in the pregnant animal and her offspring (Matzuk et al., 1998). Differential sensitivity to ethanol-induced toxicity has been demonstrated in embryonic neural crest cells from different mouse strains, presumably due to different genotypes (Chen et al., 2000), which may illustrate the interplay between (maternal) environment and fetal genotype for the induction of maldevelopment.
The WT and TG fetuses are C57/Bl/6 derived, whereas the KO mice were constructed on a 129 background, which is related to, but not identical to, the C57/Bl/6 background. However, the CKO and CTG groups did not display any differences in litter size, fetal outcome, and fetal/placental weight; therefore, we regard comparisons of the outcome of WT fetuses in all ethanol-exposed groups to be scientifically justified. The WT embryos were damaged (resorbed) more than heterozygous and homozygous embryos in the EKO animals. This finding was not expected but may be related to the fact that CuZnSOD is needed for normal ovarian function since mice that are homozygous for the targeted mutation of CuZnSOD display reduced fertility and altered ovarian morphology (Matzuk et al., 1998). However, why a general decrease in fertility may specifically affect the viability of WT offspring (compared with the viability of heterozygous and homozygous KO offspring) is not completely clear.
The finding of an unequal distribution of fetal damage in the uterine horn of the WT and KO pregnant mice, where the extreme positions of the horns display increased proportions of fetuses with developmental defects (resorptions), may indicate that the existing gradient of nutrients and ethanol (with higher levels of nutrients and ethanol towards the ends of the horn) is important for the risk of developing ethanol embryopathy, at least when the maternal antioxidative defense is not sufficient to block adverse outcome. The importance of increased nutritional supply, as a function of uterine position, has been documented before, in experimental diabetic pregnancy (Wentzel et al., 1995).
The ethanol-exposed fetuses displayed increased hepatic levels of the isoprostane 8-epi-PGF2, indicative of an involvement of oxidative stress in the etiology of the embryopathy in our mouse model. This notion has been suggested before, based on several experimental studies, in particular the idea of ethanol metabolism as a ROS producing activity has been proposed (Henderson et al., 1999; Kotch et al., 1995). The possibility that ethanol consumption leads to depletion of the cellular stores of reduced glutathione (GSH) has been shown by several research groups (Addolorato et al., 1997; Devi et al., 1993; Reyes et al., 1993). The major GSH loss may actually take place in the mitochondrion (Ramachandran et al., 2003), which therefore should both suffer from oxidative damage (Devi et al., 1994) and produce enhanced amounts of ROS (Siler-Marsiglio et al., 2005). Since enhanced susceptibility to ROS may also be present in embryonic tissues with developmental significance (Chen et al., 2004; Devi et al., 1996; Dreosti et al., 1981; Lee et al., 2005), e.g., neural crest cells (Chen et al., 2000; Davis et al., 1990), the exposure to ethanol can have teratogenic effects (Cartwright and Smith, 1995), even at low concentrations (Vaglenova and Petkov, 1998).
The presence of a severe state of ethanol-induced oxidative stress in the WT and KO animals concomitant with increased developmental damage in the offspring contrasted with the limited effect caused by ethanol in the offspring of the TG animals. This discrepancy suggests the presence of a teratological signaling system from mother to offspring in the WT and KO mice, a signaling system not present in the TG animals. A transfer of maternal substances with teratogenic properties may be an evident possibility as the teratogenic signal, substances that would be specifically enhanced by a state of oxidative stress. Biochemical candidates for such an inductive transfer would be lipid peroxides, for instance isoprostanes, which are formed in the maternal compartment and may traverse the yolk sac placenta and damage the embryo. Experimental support for the teratogenicity of isoprostanes has been obtained in vitro where rat embryos subjected to increased levels of isoprostane 8-epi-PGF2 displayed marked dysmorphogenesis (Wentzel and Eriksson, 2002). In addition, simultaneous supplementation of the antioxidative compounds N-acetylcysteine or SOD effectively blocked the isoprostane-induced damage to the embryos, thereby suggesting that the metabolism of the product of oxidative stress, 8-epi-PGF2, may indeed promote further ROS production in the embryonic tissues (Wentzel and Eriksson, 2002). Alternatively, the isoprostanes may exert other metabolic and cellular effects in the embryo since they have both vasoconstrictor (Morrow et al., 1992; Takahashi et al., 1992) and apoptotic-cytotoxic (Brault et al., 2003) activity.
The precise nature of the teratogenic effect exerted by the maternal oxidative stress is elusive at present. The etiology of the malformations found in the present work is thus not clear; however, our findings indicate that ethanol exposure disturbs embryogenesis by enhanced maternal and embryonic oxidative stress, and the adverse effects can be partly ameliorated by enhanced antioxidative capacity in the intrauterine environment.
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ACKNOWLEDGMENTS |
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The authors are grateful to Professor Ulf Rydberg for constructive and fruitful discussions. The study was generously supported by the Swedish Labor Market Insurance Company and The Swedish Research Council (grant nos 12X-7475 and 12X-109).
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