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Abnormal placentation is a potential mechanism to explain the increased incidence of low birthweight observed after IVF. This study evaluates, in a mouse model, whether the method of conception and embryo transfer affect placentation and fetal development.
IVF blastocysts (CF1 × B6D2F1/J) were cultured in Whitten's medium (IVFWM, n = 55) or K modified simplex optimized medium with amino acids (IVFKAA, n = 56). Embryos were transferred to the uteri of pseudo-pregnant recipients. Two control groups were created: unmanipulated embryos produced by natural mating (in vivo group, n = 64) and embryos produced by natural mating that were flushed from uterus and immediately transferred to pseudo-pregnant recipients (flushed blastocysts, FB group, n = 57). At gestation age 12.5 days, implantation sites were collected and fixed; fetuses and placentas were weighed and their developmental stage (DS) evaluated. Placental areas and vascular volume fractions were calculated; parametric statistics were applied as appropriate.
IVF fetuses showed a modest but significant delay in development compared with FB mice (P < 0.05). In addition, IVF conceptuses were consistently smaller than FB (P < 0.05). Importantly, these differences persisted when analyzing fetuses of similar DS. The placenta/fetus ratio was larger in the IVF group (IVFWM 0.95; IVFKAA = 0.90) than the FB group (0.72) (P < 0.05 for all comparisons). Gross morphology of the placenta and ratio labyrinth/fetal area were equivalent in the IVF and FB groups, as were percentage of fetal blood vessels, maternal blood spaces and trophoblastic components.
In vitro embryo culture affects fetal and placental development; this could explain the lower birthweight in IVF offspring.
Assisted reproduction techniques (ART), such as IVF and ICSI are very successful in providing excellent pregnancy rates. However, new evidence suggests that these procedures are not free of risks. Human studies show an increased incidence of maternal complications (such as pre-eclampsia and gestational diabetes) after ART (Jackson et al., 2004). Increased fetal complications, including low and very-low birthweights (Schieve et al., 2002), imprinting disorders (Ludwig et al., 2005; Sutcliffe et al., 2006) congenital malformations (Hansen et al., 2002), and potentially long-term effects on offspring health (Ceelen et al., 2007, 2008a, b, c) have been reported. Data from animal models reveal that blastocysts produced by ART show gene expression changes (Giritharan et al., 2007) and abnormal imprinting marks (Doherty et al., 2000); ART mice offspring have been shown to exhibit abnormal behavior (Ecker et al., 2004; Fernandez-Gonzalez et al., 2004) or an altogether abnormal phenotype (such as the large offspring syndrome in cattle; Young et al., 1998).
In addition, pregnancies resulting from ART show an increased incidence of anomalous placentation, including abnormal shape, abnormal umbilical cord insertion and presence of vasa previa (Gavriil et al., 1993; Al-Khaduri et al., 2007). A 3–5-fold increased risk of placenta previa has been associated with ART (Jackson et al., 2004; Romundstad et al., 2006). Studies in cattle show abnormal gene expression and a reduction in placentome number following somatic cell nuclear transfer (Arnold et al., 2006; Everts et al., 2008). The number of trophectodermal cells is reduced in IVF blastocysts; in addition genes involved in placental transport function are misregulated after IVF (Giritharan et al., 2007).
The etiology of these complications is likely diverse: the infertile population could be predisposed to diseases (Draper et al., 1999; Bahceci et al., 2005; Kapiteijn et al., 2006); epigenetic dysregulation, or altered placentation following in vitro culture are additional possibilities. Given the key role of the placenta throughout pregnancy, and the evidence of placental abnormalities in ART pregnancies, we hypothesized that ART may lead to suboptimal placentation that may cause impaired embryo development. Compromised placental structure or function may be a cause or contributory factor in obstetrical and neonatal complications associated with assisted reproduction. To test if different culture conditions affect embryo development and placentation, we initially cultured embryos in suboptimal conditions, using Whitten's medium (WM and 20% O2). Whenever an effect on placental or embryonic growth was noted using WM, the experiment was repeated using more optimal culture conditions, with K modified simplex optimized medium + amino acids (KSOM + AA) under 5% O2 (Rinaudo and Schultz, 2004).
IVF was performed as previously described (Giritharan et al., 2007). Briefly, 6 weeks old CF1 female mice were injected with 5 IU pregnant mare's serum gonadotrophin (PMSG) and 42–46 h later with 5 IU hCG. Oocytes were collected from the ampullae 13 h after hCG injection. Sperm was obtained from the cauda epididymis of B6D2F1/J males.
Gametes were co-incubated in WM containing 15 mg/ml bovine serum albumin (BSA) for 4 h. Fertilized oocytes were washed and cultured to the blastocyst stage in WM under mineral oil in a 37°C humidified atmosphere with either 20% O2 or 5% O2 in modular incubators (IVFWM group).
For those experimental conditions where differences between IVFWM and control mice were noted, an additional IVF group was created using optimized culture conditions (KSOM + AA under 5% O2, the IVFKAA group) as a way to assess if culture conditions alone were responsible for the outcome.
Naturally ovulating female CF1 mice, mated with vasectomized CD1 males, were used as recipients for embryo transfers. Mating was confirmed by the presence of a vaginal plug the following morning. Blastocyst transfers were performed 2.5 days post coitum. Late-cavitating blastocysts of similar morphology were then transferred to the uterus (n = 10–12 per horn) of pseudo-pregnant recipients. It is important to note that the same number of embryos was transferred in all groups [IVF and flushed blastocysts (FB)] and therefore no bias was introduced because of this technical aspect.
Control mice were generated as follows:
The protocol for animal handling and treatment procedures was reviewed and approved by the Animal Care Facility at University of California San Francisco.
Gestational age (GA) for the in vivo group was calculated by considering a positive plug as Day 0.5 of pregnancy, whereas GA for embryos undergoing embryo transfer was calculated according to the state of pseudo pregnancy of the surrogate mother (Van der Auwera and D'Hooghe, 2001; Nagy and Vintersten, 2003); specifically, the day of transfer was considered Day 2 of gestation (Supplementary data, Fig. S1 for detailed explanation).
At 12.5 days GA in the afternoon (3 p.m. ± 1 h), pregnant mothers were euthanized by CO2 inhalation and cervical dislocation and fetuses and placentas were recovered. All implantation sites were counted and examined and designated as viable or abortive. Abortive sites were defined as the sites at which an implantation site was detectable but the embryo was not identifiable (because necrotic or not vascularized). Only implantation sites where both placenta and fetuses could be assessed were considered viable.
Dissection of the implantation sites was performed under an Olympus SZX9 dissecting microscope. All the implantation sites in the FB (57 fetuses from 5 litters), IVFWM (55 fetuses from 5 litters), IVFKAA (56 fetuses and 5 litters) and in vivo groups (64 fetuses from 5 litters) could be examined.
Abortion rate was defined as the ratio of abortive sites/total implantation sites. After dissection, each fetus and placenta at 12.5 days GA was immediately fixed overnight in 4% paraformaldehyde. After 24 h, fetuses and placentas were weighed individually, and stored in 70% ethanol. Gestational sacs containing more than one fetus were not used for statistical analysis.
We defined embryonic developmental stage (DS) using surface anatomy, according to the method of Gruneberg (1943).
Fixed placental discs were bisected through the attachment of the umbilical cord and embedded in paraffin wax. At least seven placentas (from different litters) in each group were examined. Fixed placentas were weighed (wet weight, g) and sampled. Six micrometer cross sections were obtained and stained with hematoxylin and eosin. The resulting sections were observed and photographed with a Leica microscope and camera (Model DMRB, Leica Microsystems AG, Wetzlar, Germany). The border between the labyrinth and spongiotrophoblast was identified visually and outlined using Photoshop software (Adobe, San Jose, CA, USA). The border between maternal and fetal components of the placenta was identified by the presence of trophoblast giant cells; a continuous line connecting the trophoblast giant cells was created likewise using Photoshop. This method of standardization allowed us to control for the variable collection of the decidual component of the placenta. The total cross-sectional areas of fetal and labyrinthine zones were measured and calculated using the Photoshop plug-ins Image Processing Toolkit (Reindeer Graphics, Asheville, NC, USA).
To evaluate if proliferation or distribution of maternal blood spaces, fetal blood vessels and trophoblast cells differed among groups, a subgroup of placentas was stained, using Ki67 for proliferation and MTS-12 for endothelial cells. Briefly, six micron paraffin sections were deparaffinized in Citrisolve, rehydrated to phosphate-buffered saline (PBS) and treated with 0.3% hydrogen peroxide to block endogenous peroxidase. The sections were treated with blocking solution (1%BSA/PBS). For proliferation analysis, slides were incubated in boiling unmasking solution (Vector, Burlingame, CA, USA) for 30 min, cooled and exposed to a 1:10 dilution of Ki67 antibody (NCL-Ki67-MM1; Novacastra) then treated with a secondary antibody conjugated to polymer-horse-radish peroxidase (Dako Carpinteria, CA, USA).
For endothelial staining, slides were treated with pronase (0.01%, Sigma), washed and incubated in undiluted rat monoclonal MTS-12 (generous gift of Dr Richard Boyd, Monash University), then treated with biotinylated anti-rat antibody and ABC reagent (Vector). Diaminobenzidine and Meyer's hematoxylin (both from Sigma, St. Louis, MO, USA) were used as chromagen and counter stain, respectively.
The percentage of different placental components (fetal capillaries, trophoblast and maternal blood spaces) in a field of view was estimated using imageJ software (version 1.42q, National institute of Health, USA; Bergmann et al., 2004; McArdle et al., 2009). Fields of view randomly sampled from different placentas (n = 7 IVF; n = 5 FB) were photographed using a 40× objective lens. The percentage of proliferating cells was estimated by counting ~1300 cells per animal (n = 7 IVF; n = 8 FB).
All data are presented as mean ± SD; one way analysis of variance was used for statistical analysis as appropriate; Bonferroni post hoc correction was applied if P < 0.05. P-value of <0.05 was considered significant.
The total number of implantation sites was similar in all groups. However, IVFWM mice showed a significantly higher abortion rate (26.89%, P < 0.05) when compared with FB (7.28%) or in vivo animals (7.09%, Table I). The abortion rate was not significantly different between embryos cultured in WM or KSOM, or between IVFKAA (17.00%) and control groups. The twinning rate was 0% in FB, 1% in in vivo, 0% in IVFKAA and 5.4% in IVFWM group.
The fact that IVF and FB fetuses have similar numbers of implantation sites is important because litter size can affect growth parameters. Indeed superovulated in vivo mothers have more implantation sites, larger litter sizes, higher abortion rate and smaller fetuses than in vivo non-superovulated mothers (Supplementary data, Table SI).
Litters displayed significant variability of development and weight. The sex ratio was not different among the groups. Overall, at 12.5 days GA IVFWM fetuses (64.4 ± 15.7 mg) and placentas (58.1 ± 12.7 mg) were significantly smaller than those in the FB group (98.7 ± 26.5 and 68.0 ± 16.2 mg, respectively; Table II); IVFKAA fetuses weighed less (79.18 ± 0.5 mg) than FB fetuses but more than IVFWM fetuses. Placental weight in the IVFKAA group (69.47 ± 0.5 mg) was similar to the placentas of FB fetuses and were larger than IVFWM (Table II and Fig. 1).
The placenta/fetus ratio was significantly larger in the IVF groups (0.90 for IVFKAA and 0.95 for IVFWM) than in the FB group (0.72, P < 0.05). Of note the FB group had a ratio similar to the in vivo group (0.68).
Because IVF embryos reach the blastocyst stage with several hours of delay compared with in vivo embryos (Giritharan et al., 2007), we investigated whether this delay in development was maintained for the remainder of gestation. At 12.5 days GA IVFWM fetuses were slightly delayed compared with FB fetuses (FB = 12.80 ± 0.45 days; IVF–WM = 12.42 ± 0.47 days). IVFKAA fetuses (12.62 ± 0.53) displayed an intermediate delay (Fig. 1C), suggesting a graded effect of the in vitro culture conditions on development.
To control for the fact that fetuses that have a delayed DS might have a reduced weight, we analyzed a subgroup of fetuses at a similar DS. Importantly, the weight differences persisted when analyzing fetuses at a similar DS (13 days, Table II).
The findings of smaller placentas and a larger placenta/fetus ratio in IVF fetuses than FB, prompted us to analyze the histological characteristics of the placenta. We compared the placenta from animals with the same DS (13 days GA).
Gross placenta morphology did not differ in the three experimental groups: labyrinth, spongiotrophoblast development and trophoblast giant cells appeared to be similar when analyzed in cross sections through the umbilical vessels insertion (Fig. 2). The total placental area (7.5 ± 1.6 mm2) at 13 days GA was larger in FB than in vivo (5.3 ± 0.7 mm2) and IVFWM placentas (5.5 ± 1.5 mm2), P < 0.05, which is in accordance with the weight data. Nevertheless the ratio of labyrinth area and spongiotrophoblast area was similar in the three study groups (0.69 for in vivo; 0.66 for FB and 0.63 for IVFWM).
Analysis of placenta cell proliferation, measured with Ki67 antibodies, revealed that IVFWM placenta had slightly increased proliferation compared with FB (Table III). Analysis of the volume of fetal blood vessels, maternal blood spaces and trophoblast compartments did not show differences between IVFWM and FB placentas.
We provide evidence that IVF and embryo culture affect the in utero fetal and placental development of CF1 × B6D2F1/J mice. IVF fetuses display a modest but significant delay in development and are smaller than FB fetuses, also after correcting for DS. Furthermore, there is a more obvious reduction in placental and fetal weights as the preimplantation stress increases (IVFWM weigh less than IVFKAA). These findings suggest that suboptimal placentation follows embryo culture in vitro and could explain the lower birthweight observed in offspring conceived in vitro.
A number of important conclusions can be derived from these experiments. Although embryos undergoing only the embryo transfer procedure (FB group) adapt their development to the surrogate GA, the fetuses generated in vitro and transferred to recipient mothers (IVF group) show a more noticeable phenotype: their observed development is lagging behind their expected development (by 0.38 days); it is interesting to note that IVF conceptuses maintained the delay in development that was accrued after culture at the blastocyst stage (and showed neither a ‘catch up’ growth nor a magnified delay in development). IVF fetuses and placentas are smaller than FB fetuses even when controlling for DS (Fig. 1 and Table II). An improved media (KAA) partially compensates for the effect of culture; however, even optimized in vitro culture conditions affect fetal development (Fig. 1).
The placental/fetus ratio progressively and significantly increased as conditions deviated further from the in vivo situation. The placenta/fetus ratio is considered a marker of intrauterine stress (Barker et al., 1990). Whereas both fetal and placental weights decrease as a result of manipulation in vitro, there is a greater decrease in the size of the embryo than in placenta size (Fig. 1).
The reasons for the increase in placental/fetus ratio could be diverse and a combination of different factors may be involved. It is important to note that an increased placenta/fetus ratio has been described in human IVF pregnancies and has been associated with hypertension later in life (Barker et al., 1990; Moore et al., 1996; Koudstaal et al., 2000).
These effects cannot be attributed to differences in the number of embryos transferred, or to differences in implantation rate or litter size. In fact, a similar number of embryos were transferred in FB and IVF groups and a similar number of embryos implanted (Table I), allowing a consistent comparison between groups. However, there is a higher incidence of abortive sites in IVF mice when compared with FB animals. Interestingly there was a higher twinning rate in IVFWM fetuses: although the increase was not significant, it is worth noting, since twinning rate is very rare in naturally mated mice. It remains to be evaluated if this effect is induced by ovulation induction or embryo culture.
Because of the experimental design, we do not know if placental size and ratio will be maintained at birth. The altered placenta/fetus ratio was not associated with an abnormal placental morphology. IVFWM, in vivo and FB placentas displayed a similar labyrinth and spongiotrophoblast development. The total placenta and labyrinth areas were larger in the FB group compared with in vivo and IVFWM group but the ratio labyrinth/total area was similar, suggesting normal function. It has been noted that changes in the labyrinth/total area ratio are associated with impaired placenta function (Rossant and Cross, 2001). Because histological results were similar in IVFWM and FB groups, we did not repeat the experiments with IVFKAA group. Placental fetal blood vessels, maternal blood spaces and trophoblast compartments were similar in FB and IVFWM. Only Ki67 staining was increased in 12.5 day GA IVF placentas (P < 0.05). We have evidence (data not shown) that placenta morphology and proliferation is not different at GA Day 8.5 in FB and IVFWM groups. It is possible that an increase in mitosis at GA 12.5 days represents a compensatory mechanism to rescue an originally restricted placental tissue mass. However, because Ki67 is a marker both of proliferation and endoreduplication (typical of trophoblast giant cells) our findings do not necessarily indicate an increase in proliferation and placental growth.
Placental size can be affected by gamete manipulation or embryo culture. In cattle, large offspring syndrome is associated with normal placental size (Sinclair et al., 1999) and interestingly, the gene expression profile of the endometrium of IVF and in vivo generated cows is different, suggesting that the endometrium functions as a biological sensor of in vitro manipulation technologies (Mansouri-Attia et al., 2009).
It has yet to be elucidated whether IVF alters placental function. Collier reported dysregulation of placental steroid metabolism in mouse pregnancies conceived through ART (Collier et al., 2009). Proteomic analysis reveals the presence of an abnormal protein profile after ART (Zhang et al., 2008). Our results support epidemiologic data, indicating that IVF term singletons weigh less than naturally conceived children (McDonald et al., 2009). These findings also validate blastocyst gene expression data, which show that in vitro embryos have an altered gene expression pattern with increase in apoptotic and ‘stress’ pathways (Rinaudo and Schultz, 2004; Rinaudo et al., 2006; Giritharan et al., 2007). Our laboratory has shown that IVF blastocysts contain fewer trophectodermal cells (Giritharan et al., 2007) which may lead to a smaller placenta, resulting in an even smaller fetus.
The mechanism by which IVF affects placental or fetal size is unknown. One possibility is that epigenetic differences are responsible for these findings. Indeed Rivera et al. (2008) show that the process of embryo transfer alone is sufficient to alter the methylation pattern of several imprinted genes. Perturbations in embryonic metabolism and gene expression may be additional mechanisms through which in vitro culture influences embryonic viability and subsequent development.
It remains to be determined whether these perturbations of fetal growth and development will be associated with longer-term health consequences.
These findings need further investigation since they suggest a possible suboptimal pattern of placentation in IVF that differs from that of an in vivo-conceived control group. If a similar impairment occurs in humans, it may explain the obstetric and perinatal complications associated with ART.
Several considerations need to be kept in mind when analyzing the current study. First, differences between human and mouse placentation need to be taken into account. Mouse and human placenta differ in their morphogenesis and exchange functions (Rossant and Cross, 2001; Malassine et al., 2003). On the other hand, placentation is hemochorial in both species and similarities have been identified among placental cell populations (Malassine et al., 2003; Wang and Dey, 2006).
Second, the decision to sacrifice mice at 12.5 days GA was based on developmental considerations. Between 8 and 10 days post-fertilization the placenta has completed the fusion of chorion and allantois and can be defined as mature (Rossant and Cross, 2001). From this stage onwards the placental architecture is well established and spongiotrophoblast and labyrinthine zones can be distinguished clearly. Although additional time points could have been selected to observe placenta and embryo development, analysis of placenta on Day 12.5 offers sufficient sensitivity to describe a delay in placenta maturation and morphology.
Third, we used morphologic characteristics to define DS, following the protocol of Gruneberg (1943) which allows the dating of mouse fetuses from 7 to 18 days with an error of <24 h, based on the external features of the fetus. If fetuses had developmental characteristics that were between 10 and 11 days of development they were given an intermediate score (a value of 10.5 days). Alternative methods, such as somite counting or crown rump length (CRL) measurements, have an even higher margin of error. In fact, by embryo Day 10 most of the rostral somites already begin to de-epithelialize into sclerotomal cells and become difficult to count (Kei Chan et al., 2005). CRL, on the other hand, is an unreliable estimator of embryonic age, since recoil and stretching of the embryo itself can fix it in abnormal postures. It is possible that the Gruneberg method we utilized also lacks precision. However, two independent observers, blinded on the study group, analyzed all the data and reached the same conclusions. Furthermore, the in vivo group showed a close correspondence between the observed and expected development.
In summary, we describe for the first time the effect of the method of fertilization on placentation, controlling for the effect of the embryo transfer procedure. We show that placental and fetal weights, as well as the placenta/fetus ratios, are affected by preimplantation embryo manipulation.
L.D.P.: collection and assembly of data and manuscript writing. W.L.: collection of data. X.L.: collection of data. A.D.: collection of data and manuscript writing. P.M.: collection of data. A.R.: data analysis and interpretation. E.M.: conception and design, data analysis and interpretation. P.R.: conception and design, data analysis and interpretation manuscript writing and final approval of manuscript.
This research was supported by NICHD/NIH through cooperative agreement 1U54HD055764 as part of the Specialized Cooperative Centers Program in Reproduction and Infertility Research.
The author would like to thank Dr Marco Conti for the helpful suggestions on reading the manuscript. MTS-12 rat monoclonal antibody was generously donated by Dr Richard Boyd, Monash University, Australia.