Abstract
Iron deficiency, which occurs when iron demands chronically exceed intake, is prevalent in pregnant women. Iron deficiency during pregnancy poses major risks for the baby, including fetal growth restriction and long-term health complications. The placenta serves as the interface between a pregnant mother and her baby, and it ensures adequate nutrient provisions for the fetus. Thus, maternal iron deficiency may impact fetal growth and development by altering placental function. We used a rat model of diet-induced iron deficiency to investigate changes in placental growth and development. Pregnant Sprague-Dawley rats were fed either a low-iron or iron-replete diet starting 2 weeks before mating. Compared with controls, both maternal and fetal hemoglobin were reduced in dams fed low-iron diets. Iron deficiency decreased fetal liver and body weight, but not brain, heart, or kidney weight. Placental weight was increased in iron deficiency, due primarily to expansion of the placental junctional zone. The stimulatory effect of iron deficiency on junctional zone development was recapitulated in vitro, as exposure of rat trophoblast stem cells to the iron chelator deferoxamine increased differentiation toward junctional zone trophoblast subtypes. Gene expression analysis revealed 464 transcripts changed at least 1.5-fold (P < 0.05) in placentas from iron-deficient dams, including altered expression of genes associated with oxygen transport and lipoprotein metabolism. Expression of genes associated with iron homeostasis was unchanged despite differences in levels of their encoded proteins. Our findings reveal robust changes in placentation during maternal iron deficiency, which could contribute to the increased risk of fetal distress in these pregnancies.
Keywords: iron deficiency, nutrient, placenta, anemia, trophoblast, pregnancy
Iron deficiency is the most common nutrient insufficiency worldwide, affecting more than 20% of the global population (1). It occurs after chronic imbalance between iron intake and demand, leading to exhaustion of tissue iron stores. Due to the necessity of iron for hemoglobin (Hb) synthesis, iron deficiency can progress to anemia: a condition in which the number of red blood cells falls below clinical thresholds (2). Since iron is essential to support placental and fetal growth and to maintain red blood cell mass during maternal blood volume expansion, pregnant women are at high risk of iron deficiency and anemia. Approximately 38% of women develop anemia during pregnancy, including 22% of women in developed countries, with most cases attributed to iron deficiency (3, 4).
Maternal iron deficiency can have major repercussions on fetal health. Iron deficiency during pregnancy increases the risk of fetal death, low birth weight, and preterm birth, and it is associated with long-term cardiovascular, metabolic, neurobehavioral, and immunological complications in affected offspring (5). Despite repletion of iron stores in children receiving iron supplements, health complications persist in children if mothers exhibited iron deficiency during pregnancy, emphasizing the importance of adequate iron provision in utero for long-term health of offspring (6). In an effort to prevent anemia and potential adverse health outcomes, iron supplementation is recommended for pregnant women. However, iron supplementation is rife with complexity, including poor intestinal absorption of iron, gastrointestinal side effects that diminish patient adherence, and potential toxicity to the mother and fetus (7). Furthermore, while iron supplementation is effective in treating maternal anemia, it is less clear whether pregnancy outcomes are improved following iron supplementation (8). Therefore, there is a need to uncover mechanisms through which maternal iron deficiency impacts fetal growth and development to implement more effective treatment options. As the interface between maternal and fetal circulations, the response of the placenta to maternal iron deficiency may be a key consideration.
The placenta is an extraembryonic organ that supports fetal growth and development by controlling nutrient, respiratory gas, and waste transport. It is also responsible for synthesizing and metabolizing hormones crucial for maintaining pregnancy and prompting maternal physiological adaptations, and it dampens potentially harmful maternal immune reactivity. As an adaptive organ, the placenta undergoes structural and functional changes throughout pregnancy in response to fetal, maternal and environmental stimuli. Such changes include alterations in placental size, surface area available for exchange, placental vascularization, and hormone metabolism (9). Underlying many of these structural changes are alterations in differentiation and function of specialized subtypes of trophoblasts, the parenchymal cells of the placenta. Trophoblasts arise through a multilineage differentiation pathway from multipotent trophoblast stem (TS) cells. In human placentae, trophoblasts specialize into villous trophoblasts, which line chorionic villi and regulate nutrient and gas exchange between maternal and fetal blood; or extravillous trophoblasts, that interface maternal decidua and facilitate nutrient flow to the intervillous spaces. A similar dichotomous organization exists in rodent placentae, in which trophoblasts contribute either to the labyrinth zone (specializing in nutrient and gas exchange) or junctional zone (specializing in hormone production and directing nutrient flow to the labyrinth zone).
Given that the placenta is an adaptable, metabolically active organ that promotes a consistent stream of nutrients and oxygen to the fetal-placental unit, poor iron availability may have a profound effect on placental structure and function (reviewed in (10)). For instance, several studies using large cohorts of human pregnancies have revealed an inverse relationship between placental weight and maternal Hb concentration, suggesting that the placenta may respond to low iron and/or Hb levels by increasing size to enhance oxygen and nutrient exchange (11, 12). Placental weights and placental:fetal weight ratios are accordingly increased during maternal anemia (11, 13-17), albeit some studies have found no correlation between low Hb and placental weight (18), or even decreased placental weights in anemic mothers (19-21). Multiple factors may account for these discrepancies, including number of samples analyzed, ethnic populations, severity of anemia, timing of anemia onset, mode of delivery, and possibility of maternal malnutrition or other nutritional deficits. Studies using rodents fed iron-deficient diets during pregnancy, which are advantageous to control environmental and experimental variables, have typically identified growth-restricted fetuses/neonates (22-26) associated with increased placental weight or placental:fetal weight ratios (23-25, 27, 28). In general, these findings in rodents are consistent with studies using human placentas, and could reveal a conserved compensatory or pathological response of placental development to maternal iron deficiency.
In this study, we used our well-established model of maternal iron restriction in pregnant rats (24, 25, 29, 30), which recapitulates the gradual depletion of tissue iron stores and reduced fetal growth characteristic of maternal anemia in humans, to investigate how maternal iron deficiency affects placental structure. We found that placentas were larger in dams fed low-iron diets, due primarily to expansion of the junctional zone. These effects were consistent in vitro, where TS cells cultured with an iron chelator are primed to form junctional zone lineages. Furthermore, gene expression profiling revealed changes associated with oxygen and nutrient transport, lipoprotein metabolism, hormone responses, and cell communication in iron-deficient placentas. Our findings suggest that maternal iron deficiency has profound effects on placentation, which could contribute to poor pregnancy health and adverse outcomes in offspring.
Methods
Animals and Treatments
All protocols involving animals were approved by the University of Alberta Animal Care Committee in accordance with guidelines established by the Canadian Council for Animal Care. Sprague-Dawley rats (aged 6 weeks) were purchased from Charles River (Saint-Constant, QC, Canada) and housed at the University of Alberta animal care facility under a 12-hour light/dark cycle and an ambient temperature of 23 °C. Dams had ad libitum access to food and water throughout the study.
All purified diets used in this study were based on the AIN-93G formula (Research Diets, New Brunswick, NJ), which contained varying amounts of iron in the form of ferric citrate. Two weeks before mating, rats were randomly assigned either to a control diet group (37 mg/kg iron diet; D10012G) or an iron-restricted diet group (3 mg/kg iron diet; D03072501) to elicit depletion of tissue iron stores. After 2 weeks on their respective diets, female rats were bred to age-matched males that were fed a standard rodent chow (5L0D; PicoLab, St. Louis, MO). The presence of sperm in a vaginal smear the next morning was used to confirm pregnancy and was considered gestational day (GD) 0.5. Dams were single-housed thereafter. Sperm was detected in 17 dams fed the control diet, and 18 dams fed the iron-deficient diet. Dams in the control group were maintained on the same control diet throughout pregnancy. Dams that were fed the iron-restricted diet before pregnancy were transferred to a diet that contained 10 mg/kg iron (D15092501) for the duration of the study, as previously described (24, 25, 29). Throughout pregnancy, maternal food consumption, body weight, and Hb concentration were measured weekly. Maternal Hb levels were assessed from blood (~10 μL) collected by saphenous venipuncture, analyzed on a HemoCue 201+ system (HemoCue, Ängelholm, Sweden).
Rats were anesthetized on GD13.5 (control: n = 9, iron deficiency: n = 8) or GD18.5 (control: n = 8, iron deficiency: n = 10) with isoflurane (5% induction, 3% maintenance in pure O2), then euthanized by exsanguination. Conceptuses were dissected, and fetuses and their placentae were subsequently weighed. The mean litter size among all groups was 14. One litter from the GD13.5 iron-deficient cohort had only 2 conceptuses (both viable), and 1 litter from the GD18.5 iron-deficient cohort had only 5 conceptuses (2 viable). These 2 pregnancies were included in maternal assessments but were excluded from experiments involving fetal and placental analyses due to the shortage of tissue. For conceptuses isolated on GD18.5, approximately half of the fetuses from each litter were decapitated, a core blood sample was obtained to assess Hb levels, and placentas and selected organs were dissected and weighed. Liver tissue was further processed to assess levels of ferritin. Placental tissue was stored at −80 °C until further processing. For conceptuses where placental morphology was investigated, whole conceptuses were placed in dry ice-cooled heptane and stored at −80 °C until cryosectioned.
Enzyme Immunoassay
Levels of ferritin in fetal liver on GD18.5 were assessed using a colorimetric enzyme immunoassay (MBS700510, RRID:AB_2893344, measures ferritin light chain; MyBioSource, San Diego, CA), performed according to the manufacturer’s instructions. Briefly, fetal livers were dissected on GD18.5, weighed, homogenized in phosphate-buffered saline (PBS), subjected to 2 consecutive freeze-thaws to facilitate tissue breakdown, centrifuged, and then the supernatant added to the coated microplate. A standard curve with no logarithmic transformation was generated using absorbance values plotted against defined concentrations of recombinant rat ferritin. Samples were diluted in assay diluent buffer to ensure absorbance values fell within the linear range of the standard curve. The sensitivity of the immunoassay for ferritin detection was 0.23 ng/mL.
Immunohistochemistry and Placental Morphology Assessments
To assess tissue morphology, GD13.5 and GD18.5 placentas were cryopreserved in Tissue-Tek 4583 optimal cutting temperature compound (ThermoFisher Scientific, Whitby, ON, Canada) and cryosectioned at 10 μm thickness. Sections were aligned vertically through the center of the placenta as indicated by the presence of a maternal arterial channel traversing into the junctional zone. For hematoxylin and eosin staining, placental cryosections were fixed in 10% neutral buffered formalin and incubated in Mayer’s hematoxylin solution (Sigma-Aldrich, Oakville, ON, Canada; 0.1% hematoxylin in distilled water containing 5% alum, 0.02% sodium iodate, and 0.1% sodium citrate) for 5 minutes followed by rinsing under tap water for 5 minutes. This was followed by incubation with eosin (Sigma-Aldrich) for 1 minute and rinsing under tap water for 5 minutes. The stained sections were mounted using Fluoromount-G mounting medium (SouthernBiotech, Birmingham, AL). For immunohistochemistry, placental cryosections were fixed in 10% neutral buffered formalin, permeabilized using 0.3% Triton X-100 and 1% bovine serum albumin in PBS and blocked in 10% normal goat serum (ThermoFisher Scientific). Sections were immersed in primary antibody specific for cytokeratin (catalog no. 628602, 1:400; RRID:AB_439775, BioLegend, San Diego, CA) overnight at 4 °C. After washing, sections were immersed in biotin-conjugated secondary anti-mouse IgG (Sigma-Aldrich) followed by Vectastain ABC Elite reagent (Vector Laboratories, Burlington, ON Canada), and color was developed using 3-amino-9-ethylcarbazole (AEC) red (ThermoFisher Scientific). Sections were counterstained with Gill No. 1 hematoxylin (Sigma-Aldrich) and mounted as described above. For immunofluorescence, cryosections were fixed, permeabilized, and blocked as above, then immersed in primary antibody specific for ALOX15 (119774, 1:150, RRID:AB_10901109, Abcam Inc, Toronto, ON, Canada). After washing, sections were incubated with Alexa Fluor 488-conjugated goat anti-mouse secondary antibody, nuclei counterstained in 4′,6-diamidino-2-phenylindole (DAPI, ThermoFisher Scientific), and sections mounted. All sections were imaged using a Nikon DS-Qi2 microscope. For morphology assessments, the perimeter of each placental zone was outlined using ImageJ software (31) and used to calculate cross-sectional area in arbitrary units. The mean cross-sectional area of each placental zone was calculated following 3 independent measurements of the zone perimeter. Area measurements in arbitrary units were then converted to mm using the scale bar embedded in the image as a reference. All measurements were conducted by 2 researchers who were blinded to the experimental treatments, and results between observers were averaged. For GD13.5 morphology assessments, measurements were performed using placentas from 14 dams, which after unblinding resulted in 8 dams fed iron-replete diets and 6 dams fed iron-deficient diets. GD18.5 assessments were conducted using placentas from 13 dams (5 control and 8 iron-deficient dams).
Clariom S Gene Expression Analysis
To gain mechanistic insight into how iron deficiency affects the placental transcriptome, global gene expression was analyzed in placental tissue using Clariom S gene expression arrays (ThermoFisher Scientific). RNA was extracted from GD18.5 placental tissue (control: n = 4 dams, iron deficiency: n = 4 dams, with 2 placental RNA samples combined per dam) by homogenizing in TRIzol (ThermoFisher Scientific). The aqueous phase was then diluted with 70% ethanol, placed on RNeasy columns (Qiagen Inc, Toronto, ON, Canada), treated with DNase I, and purified. RNA (25 µg per sample) was submitted to Hamilton Health Sciences Centre (Hamilton, ON, Canada) for analysis of gene expression. Expression analysis was completed on a GeneChip Scanner 3000 using the Clariom S rat array (ThermoFisher Scientific). Testing was completed using the WT Plus Reagent Kit to generate amplified and biotinylated sense-strand DNA targets in preparation for GeneChip hybridization. Using the GeneChip Fluidics Station 450, Clariom S Rat GeneChips were washed, stained, and scanned prior to hybridization and analyzed using the GeneChip Scanner 3000 and GeneAtlas Software. Data files were generated and processed with Transcriptome Analysis Console Software 4.0 (ThermoFisher Scientific) to analyze global expression patterns of genes. Data are available on the Gene Expression Omnibus (GSE163226). The Database for Annotation, Visualization and Integrated Discovery (DAVID) bioinformatics tool was used to assess biological themes in genes that exhibited altered expression in maternal iron deficiency (32).
Quantitative Reverse Transcription Polymerase Chain Reaction
Following microarray analysis, mRNA expression of selected genes was validated on independent samples by quantitative reverse transcription polymerase chain reaction (qRT-PCR). Complementary DNA was generated from purified RNA (50 ng/µL) using High Capacity cDNA Reverse Transcription kit (ThermoFisher Scientific), diluted 1:10, and used for qRT-PCR. Complementary DNA was mixed with SensiFAST SYBR green PCR Master Mix (FroggaBio, Toronto, ON, Canada) and primers described in Table 1. A CFX Connect Real-Time PCR system (Bio-Rad Laboratories, Mississauga, ON, Canada) was used for amplification and fluorescence detection. Cycling conditions were as follows: an initial holding step (95 °C for 3 minutes), followed by 40 cycles of 2-step PCR (95 °C for 10 seconds, 60 °C for 45 seconds), then a dissociation step (65 °C for 5 seconds and a sequential increase to 95 °C). Relative mRNA expression was calculated using the comparative cycle threshold (ΔΔCt) method. The geometric mean of Ct values obtained from amplification of 4 constitutively expressed genes (Actb, Gapdh, Rn18s, and Ywhaz) was used as reference RNA, as previously described (33, 34). Ct values from each of these genes were stable among the conditions tested.
Table 1.
List of primers used for qRT-PCR
Primers | Forward sequence (5′-3′) | Reverse sequence (5′-3′) | Product (bp) |
---|---|---|---|
Actb | AGCCATGTACGTAGCCATCC | CTCTCAGCTGTGGTGGTGAA | 226 |
Ak1 | GGACGATAACGAGGAGACCA | ATAGGTGCAGACCTGGGAGA | 151 |
Alox15 | CACTGTGACCTGCTGGAAAA | GGGTTAGCACCATTGAGGAA | 138 |
Apoa1 | ACAAAAACGCGAAGGAGATG | TCAGGGTAGGGTGGTTCTTG | 171 |
Cdx2 | GAGCACGGACACTGTGAGAA | AGAAGCCCCAGGAATCACTT | 201 |
Doxl1 | AGACCCATTTCCAAAGCAGA | TAAAGCGCTGCAACATGAAC | 199 |
Gapdh | AGACAGCCGCATCTTCTTGT | CTTGCCGTGGGTAGAGTCAT | 206 |
Hbz | TGCGGTTAAGAACATCGACA | CAGGACAGAAGACAGGATGGA | 205 |
Ifit1 | GGGGAACAAGATGAAGCAGA | GCAAGGCCCTGTGTAGAAGA | 145 |
Mmp9 | GTCTTCCCCTTCGTCTTCCT | CTGGACAGAAACCCCACTTC | 127 |
Mmp10 | GAAATGGTCACTGGGACC | TGCGCAGCAACCAGGAA | 51 |
Ocm2 | GAGCTGGGTCATGGTTTTGT | CTTGAGCTCATCTCCATCCA | 133 |
Pkd2l1 | CTGGACCTGGTGGTCATCTT | GAGCTGGGTCATGGTTTTGT | 234 |
Pla2g7 | TTTTTCGACTTCAGGCCATC | CCATTTCCTTTGGGGATTTT | 95 |
Prl3a1 | GGGTGCTTCAGCTGCTACTT | CATTTGCATGGTGAGGTTTG | 164 |
Prl3d1 | TTTGACTACCCTGCCTGGTC | CATTGGGTGCAAGACTTCAA | 198 |
Prl5a2 | CTCCTGGGCACTCCTGATAC | TTCACACATGGGCAAAGAGA | 699 |
Rbp4 | TCGACAAGGCTCGTTTCTCT | CCATGTCTGCACACACTTCC | 177 |
Rn18s | GCAATTATTCCCCATGAACG | GGCCTCACTAAACCATCCAA | 137 |
Tdgf1 | GGACTTGTTGCTGGGATAGG | AAGGCACAAGCTGGAGAGAG | 130 |
Tpbpa | TGGAGAGCGGAGATGAGATT | GGGACTGGCTACTGAGTTGG | 205 |
Ywhaz | TTGAGCAGAAGACGGAAGGT | CCTCAGCCAAGTAGCGGTAG | 200 |
Rat Trophoblast Stem Cell Culture
Blastocyst-derived rat TS cells were used to evaluate the effects of iron chelation on trophoblast cell behavior. Rat TS cells were generously provided by Michael Soares (University of Kansas Medical Center, Kansas City, KS). Cells were cultured in RPMI 1640 media (ThermoFisher Scientific) supplemented with 20% (v/v) fetal bovine serum (ThermoFisher Scientific), 100mM 2-ME (Sigma-Aldrich), 1mM sodium pyruvate (Sigma-Aldrich), 100 U/mL penicillin, 100 μg/mL streptomycin, fibroblast growth factor 4 (25 ng/mL; Bio-Techne, Minneapolis, MN), heparin (1 μg/mL; Sigma-Aldrich), and activin (10 ng/mL, Bio-Techne). A total of 70% of the media was preconditioned by mitomycin C-treated mouse embryonic fibroblasts prior to being added to rat TS cells, as described previously (35). To determine the effect of iron depletion on TS cell function, the iron chelator deferoxamine (25 and 50μM diluted in H2O, Sigma-Aldrich) was added to TS cells for 48 hours.
Immunofluorescence
Immunofluorescence for trophoblast-specific protein alpha (TPBPA) was conducted as previously described (36). Briefly, rat TS cells cultured in the stem state on coverslips and exposed (or not) to deferoxamine were fixed in 4% paraformaldehyde, permeabilized using 0.3% Triton X-100 and 1% bovine serum albumin in PBS, and then blocked in 10% normal goat serum (ThermoFisher Scientific). Cells were then immersed in primary antibody specific for TPBPA (104401, RRID:AB_10901888, 1:100, Abcam Inc), followed by Alexa Fluor 488-conjugated anti-rabbit secondary antibodies. Cell nuclei were counterstained using DAPI, and cells were imaged using a Nikon DS-Qi2 microscope.
Western Blotting
Western blotting was performed as previously described (37). Briefly, placental lysates were prepared by homogenizing in radioimmunoprecipitation assay buffer (50 mM Tris, 150 mM NaCl, 1% NP40, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate [SDS]) supplemented with protease inhibitor cocktail (Sigma-Aldrich). Lysates (40 μg) were mixed with 4× reducing loading buffer (0.25 M Tris, 8% SDS, 30% glycerol, 0.02% bromophenol blue, 0.3 M dithiothreitol), boiled for 5 minutes, and subjected to SDS-polyacrylamide gel electrophoresis. Proteins were transferred to polyvinylidene difluoride membranes and probed using antibodies for Ferritin Light Chain (69090, RRID:AB_1523609, 1:1000, Abcam Inc), Ferritin Heavy Chain (4393, RRID:AB_11217441, 1:1000, Cell Signaling Technology, Danvers, MA), Transferrin Receptor 1 (136800, RRID:AB_2533029, 1:2000, ThermoFisher Scientific), and GAPDH (5174, RRID:AB_10622025, Cell Signaling Technology, 1:2000). Membranes were then incubated for 1 hour with species-appropriate secondary antibodies, and signals detected using a LI-COR Odyssey imaging system (LI-COR Biosciences, Lincoln, NE).
Statistical Analyses
Values are expressed as mean ± standard error of the mean (SEM). Statistical significance was analyzed by unpaired Student’s t test when comparing 2 groups and either 1-way or 2-way analysis of variance (ANOVA) with Tukey’s multiple comparisons test when comparing 3 or more groups. All statistical tests were considered significant when P < 0.05. Statistical analyses were conducted using GraphPad Prism version 7.0 (GraphPad Software Inc.). The specific number of dams, fetuses, and placentas analyzed is indicated in the figure legends.
Results
Impact of Iron Status on Maternal and Fetal Outcomes
Our first goal was to determine the impact of feeding a low-iron diet on maternal feeding behavior and pregnancy outcome. Iron restriction prior to and throughout pregnancy had no impact on total food consumption by pregnant dams, nor did it affect maternal cumulative weight gain (Fig. 1a and 1b). Litter size and number of resorptions per litter were also not significantly different between dams fed iron-replete and iron-deficient diets (Fig. 1c and 1d). We confirmed that reduced Hb was evident in dams and fetuses fed a low iron diet, an indicator of anemia. Exposure of female rats to a low iron diet starting 2 weeks before mating caused a progressive reduction in blood Hb levels (Fig. 1e; P < 0.0001). Hb levels dipped below 10 g/dL on GD14.5 and GD18.5. By GD18.5, maternal iron deficiency resulted in a 41% decrease in both maternal Hb and fetal Hb compared with controls (Fig. 1e and 1f). Paradoxically, relative ferritin levels in fetal liver were unchanged on GD18.5 (not shown). Fetuses from iron-deficient dams had 13% lower body weights by GD18.5 (Fig. 2a), despite no differences on GD13.5. On GD18.5, iron restriction resulted in a 16% reduction in relative liver weight, but no differences in relative brain, heart, or kidney weights (Fig. 2b-2e). Thus, maternal iron deficiency was associated with reduced maternal and fetal Hb and smaller fetal size.
Figure 1.
Pregnancy outcomes, litter size, and Hb levels in dams fed iron-replete and iron-deficient diets. (a) Maternal cumulative food intake over gestation. (b) Maternal cumulative weight gain over initial body weight. (c) Litter size (mean number of live fetuses per litter on GD18.5). (d) Number of resorptions per litter. In panels (a)-(d), data are presented as mean ± SEM based on measurements from 8 control-fed and 10 iron-deficient (ID) dams sacrificed on GD18.5. (e) Maternal Hb, n ≥ 5 dams per group. (f) Fetal Hb on GD18.5 (control: n = 18 fetuses, ID: n = 25 fetuses, from at least 7 dams each group; each value represents the average of the measurements obtained per litter). Data are presented as mean ± SEM. In panel (e), P values reflect 2-way ANOVA outcomes; in all other panels, P values reflect Student’s t test outcomes. An asterisk (*) denotes statistical significance (P < 0.05).
Figure 2.
Fetal body weight and organ weight changes in dams fed iron-replete and iron-deficient diets. Dams were fed iron-replete (control) or iron-deficient (ID) diets starting 2 weeks before mating. Fetuses were collected on GD18.5. (a) Fetal body weight; (b) Relative brain weight; (c) Relative liver weight; (d) Relative kidney weight; (e) Relative heart weight. Measurements from fetuses collected from 8 control and 9 ID dams (≥ 28 control and ≥ 33 ID fetuses) were used. Data are presented as mean ± SEM, with each value representing the average of the measurements obtained per litter. P values reflect Student’s t test outcomes. An asterisk (*) denotes statistical significance (P < 0.05).
Iron Status Alters Placental Weight and Morphology
On GD13.5, there was no change in placental weight or placental:fetal weight ratio between dams fed iron-replete and iron-deficient diets (Fig. 3a and 3b). However, by GD18.5, maternal iron restriction resulted in a 10% increase in placental weight and a 32% increase in placental:fetal weight ratio compared to controls (Fig. 3c and 3d), suggesting that maternal anemia is associated with inefficient placental function. Between GD13.5 and GD18.5, total placental weight increased 4.5-fold in dams fed iron-deficient diets compared with 3.9-fold in dams fed iron-replete diets. Specialized placental zones are illustrated in the schematic in Fig. 3e, and representative images of cytokeratin-immunostained placentas from iron-replete and iron-deficiency conditions on GD18.5 are shown in Fig. 3f. There was no significant difference in the area of placental zones on GD13.5 between dams fed iron-replete or iron-deficient diets (Fig. 3g and 3h). However, by GD18.5, iron deficiency caused a 34% increase in junctional zone area compared to controls, but no difference in labyrinth zone area (Fig. 3i and 3j). Likewise, labyrinth zone and junctional zone area increased 9.2-fold and 4.9-fold between GD13.5 and GD18.5 in dams fed iron-deficient diets, compared with 6.5-fold and 3.0-fold in controls, respectively. These findings indicate that iron deficiency causes placentomegaly, and this effect is particularly notable in the junctional zone.
Figure 3.
Changes in placental weight and morphology following exposure of dams to iron-replete or iron-deficient diets. Dams were fed iron-replete (control) or iron-deficient (ID) diets starting 2 weeks before mating. Placentas and fetuses were collected on GD13.5 or GD18.5. (a) Placental weight on GD13.5; (b) Placental weight relative to fetal weight on GD13.5; (c) Placental weight on GD18.5; (d) placental weight relative to fetal weight on GD18.5. (e) Schematic representation of a GD18.5 placenta. (f) Representative images of GD18.5 placentas collected from control (left) and ID (right) dams following cytokeratin immunohistochemistry. Scale bar, 1000 µm. Cross-sectional areas were measured for GD13.5 placental (g) labyrinth and (h) junctional zones, as well as for GD18.5 placental (i) labyrinth and (j) junctional zones. Data are presented as mean ± SEM. In panels (g) and (h), data represent placental measurements from 8 control-fed dams and 6 ID dams, while in panels (i) and (j), data represent 5 control-fed dams and 8 ID dams. P values reflect Student’s t test outcomes. An asterisk (*) denotes statistical significance (P < 0.05).
Impact of Iron Deficiency on Placental Gene Expression
To determine the impact of iron deficiency on placental gene expression, Clariom S arrays were used to profile gene expression differences in GD18.5 placentas collected from iron-replete and iron-deficient diets. Out of 23 188 genes, 464 transcripts were changed at least 1.5-fold (P < 0.05) in placentas from iron-deficient dams compared with controls; 230 transcripts were upregulated and 234 transcripts were downregulated (Fig. 4a and 4b). Hierarchical clustering segregated the 8 placental tissue samples into 2 statistically significant clusters (sigclust; P < 0.05): a cluster of placentas from control dams and a cluster of placentas from iron-deficient dams (Fig. 4c). Transcripts with the greatest fold increase in placentas collected from dams fed an iron-deficient diet included those involved in immunomodulation and extracellular matrix remodeling, such as Arachidonate 15-Lipoxygenase (Alox15, 18.9-fold increase), Interferon-Induced Protein With Tetratricopeptide Repeats-1 (Ifit1, 4.2-fold increase), Matrix Metalloproteinase-10 (Mmp10; 2.6-fold increase), and Matrix Metalloproteinase-9 (Mmp9; 2.4-fold increase). Downregulated transcripts included those encoding nutrient transport and hormones, including Apolipoprotein a1 (Apoa1; 4.1-fold decrease), Prolactin Family 3, Subfamily a, Member 1 (Prl3a1; 4.75-fold decrease), and Retinol-Binding Protein-4 (Rbp4; 2.63-fold decrease). Transcripts from Clariom S analyses exhibited similar directional fold changes using qRT-PCR (Fig. 4d). A list of transcripts that showed the most pronounced changes (> 2-fold) in placentas exposed to iron deficiency is presented in Table 2.
Figure 4.
Gene expression analysis of GD18.5 placentas collected from dams fed iron-replete or iron-deficient diets. (a) Pie chart summarizing the number of transcripts unaltered, upregulated (≥1.5-fold; P < 0.05) and downregulated (≥1.5-fold; P < 0.05) in maternal iron-deficiency (ID) compared with placentas from dams fed control (iron-replete) diets. (b) Volcano plot showing the distribution of differentially expressed genes with cutoff criteria of absolute fold change ≥1.5 and P < 0.05. (c) Hierarchical clustering of the top 26 differentially expressed genes in maternal ID. (d) Microarray results were validated using qRT-PCR. Microarray fold change results are presented as means; results from qRT-PCR are presented as means ± SEM, with P values reflecting Student’s t test outcomes. An asterisk (*) denotes statistical significance (P < 0.05, n = 4 dams per group, averaged from 2 placentas per dam). (e) Representative Western blots showing protein levels of FTH1, FTL1, and TFRC in placentas from maternal ID compared with controls. GAPDH was used as a loading control. Each sample represents a placenta from a different dam. On the right, densitometric analysis of band intensity relative to GAPDH, and normalized to control placentas (red dashed line) is shown. (f) ALOX15 immunofluorescence (green) in the junctional zone of placentas collected from iron-replete (control, top panels) and ID dams (bottom panels) on GD18.5. The middle panels show nuclei, which were counterstained blue with DAPI. A merged image is shown in the right panels. Scale bar = 100 μm.
Table 2.
Genes differentially expressed in GD18.5 rat placentas during maternal iron deficiency
Gene symbol | Gene name | Fold change | P value |
---|---|---|---|
Upregulated | |||
Alox15 | arachidonate 15-lipoxygenase | 18.85 | 2.64 x 10-6 |
Tdgf1 | teratocarcinoma-derived growth factor 1 | 6.65 | 0.0001 |
Hbz | hemoglobin, zeta | 5.01 | 0.0005 |
Ifit1 | interferon-induced protein with tetratricopeptide repeats 1 | 4.23 | 0.0109 |
Doxl1 | diamine oxidase-like protein 1 | 3.06 | 0.0267 |
Prl5a2 | prolactin family 5, subfamily a, member 2 | 2.71 | 0.0250 |
Mmp10 | matrix metallopeptidase 10 | 2.55 | 0.0113 |
Pkd2l1 | polycystic kidney disease 2-like 1 | 2.54 | 0.0045 |
Mmp9 | matrix metallopeptidase 9 | 2.44 | 0.0131 |
LOC102555390; Kirrel3 | uncharacterized LOC 102555390; kin of IRRE like 3 (Drosophilia) (Kirrel3), mRNA | 2.42 | 0.0055 |
Ak1 | adenylate kinase 1 | 2.31 | 0.0013 |
Gsn | gelsolin | 2.25 | 0.0147 |
Wdr17 | WD repeat domain 17 | 2.21 | 0.0074 |
Neto2 | neuropilin (NRP) and tolloid (TLL)-like 2 | 2.21 | 0.0021 |
Fgfbp1 | fibroblast growth factor binding protein 1 | 2.16 | 0.0030 |
Dsc3 | desmocollin 3 | 2.14 | 0.0308 |
Lrp8 | low density lipoprotein receptor-related protein 8 | 2.09 | 0.0195 |
Klk15 | kallikrein related peptidase 15 | 2.08 | 0.0014 |
Vdr | vitamin D receptor | 2.01 | 0.0013 |
Gfi1b | growth factor independent 1B transcription repressor | 2.00 | 0.0022 |
Lrrc15 | leucine rich repeat containing 15 | 2.00 | 0.0052 |
Downregulated | |||
Prl3a1 | prolactin family 3, subfamily a, member 1 | -4.75 | 0.0013 |
LOC102551298 | MLV-related proviral Env polyprotein-like | -4.14 | 0.0045 |
Apoa1 | apolipoprotein A-I | -4.05 | 0.0348 |
LOC102552104 | MLV-related proviral Env polyprotein-like | -3.91 | 0.0007 |
LOC102555039 | MLV-related proviral Env polyprotein-like | -3.91 | 0.0007 |
Ocm2 | oncomodulin 2 | -3.28 | 0.0066 |
Rbp4 | retinol binding protein 4, plasma | -2.63 | 0.0306 |
Il15ra | interleukin 15 receptor, alpha | -2.57 | 0.0021 |
Pla2g7 | phospholipase A2, group VII | -2.51 | 0.0012 |
Sorbs2 | sorbin and SH3 domain containing 2 | -2.49 | 3.29 x 10-5 |
LOC102555324 | MLV-related proviral Env polyprotein-like | -2.45 | 0.0059 |
Fmo1 | flavin containing monooxygenase 1 | -2.43 | 0.0016 |
RGD1311103; Ooep | Protein RGD1311103; oocyte expressed protein | -2.37 | 0.0010 |
Fga | fibrinogen alpha chain | -2.36 | 0.0453 |
LOC102553221 | MLV-related proviral Env polyprotein-like | -2.27 | 0.0032 |
Gc | group specific component | -2.18 | 0.0143 |
Pdzk1ip1 | PDZK1 interacting protein 1 | -2.18 | 0.0035 |
Tmsbl1 | thymosin beta-like protein 1 | -2.12 | 0.0042 |
Dmkn | dermokine | -2.12 | 0.0004 |
Tmem100 | transmembrane protein 100 | -2.10 | 0.0044 |
Mlh1 | mutL homolog 1 | -2.08 | 0.0038 |
Cdc25b | cell division cycle 25B | -2.08 | 0.0156 |
Cpt1b | carnitine palmitoyltransferase 1b, muscle | -2.04 | 0.0016 |
Aadat | aminoadipate aminotransferase | -2.04 | 0.0032 |
Mmp14 | matrix metallopeptidase 14 (membrane-inserted) | -2.02 | 0.0023 |
Col1a2 | collagen, type I, alpha 2 | -2.01 | 0.0182 |
Emp3 | epithelial membrane protein 3 | -2.01 | 0.0139 |
Although we detected a change in expression of Slc11a2, which encodes Divalent Metal Transporter-1 (DMT1, 1.5-fold increase; P = 0.01), changes in expression of other genes associated with iron storage and transport were mild or not apparent (ie, no change in expression of Tfrc, which encodes Transferrin receptor (P = 0.12); Fth1, which encodes Ferritin Heavy Chain (P = 0.89); and Ftl1, which encodes Ferritin Light Chain (P = 0.59), data not shown). However, protein levels of TFRC (increased 1.6-fold), FTH1 (decreased 2.4-fold), and FTL1 (decreased 2.2-fold) were all robustly changed in placentas collected from iron-deficient dams (Fig. 4e; all P < 0.05). These findings showing robust changes in levels of proteins associated with iron homeostasis despite mild effects on expression of the genes encoding these proteins are consistent with a recent study comparing levels of iron-associated transcripts and proteins in mid-gestation mouse placenta (38), and they are not surprising given that proteins involved in iron transport are primarily regulated by posttranscriptional mechanisms (39).
Since Alox15 was the most highly upregulated transcript in placentas during maternal iron deficiency, we performed immunofluorescence to detect localization of ALOX15 in rat placenta on GD18.5. ALOX15 was detectable throughout the placenta, with staining intensity highest within junctional zone trophoblasts. In maternal iron deficiency, intensity of ALOX15 staining in the junctional zone was noticeably more intense (Fig. 4f).
Gene ontology (GO) pathway analysis comparing placentas from iron-deficient dams vs iron-replete dams showed upregulation in pathways involved in G-protein coupled receptor signaling, oxygen transport, and cell communication (Fig. 5a). Those with significant downregulation included pathways involved in lipoprotein metabolism, vitamin transport, response to estradiol, and cell adhesion (Fig. 5b).
Figure 5.
Gene ontology of placental genes differentially expressed in maternal ID. Biological processes affected by (a) upregulated genes and (b) downregulated genes in maternal iron deficiency, as determined by DAVID. The gene ontology identification number and a list of differentially expressed genes included in each category are provided.
Iron Chelation Stimulates TS Cells to Differentiate Into Junctional Zone Lineages
We sought to determine the effect of iron deficiency on TS cell function. Deferoxamine is an iron chelator and has been used to simulate an iron-deficient environment in vitro (40-43). Thus, we treated rat TS cells with various doses of deferoxamine and determined the effect on TS cell function. Doses of deferoxamine higher than 100μM caused noticeable TS cell loss of viability, so 25μM and 50μM deferoxamine were used in all subsequent experiments. First, we evaluated expression of several genes that were changed in placentas during maternal iron deficiency. Some of these genes, including Alox15 and Prl3a1, were not expressed at detectable levels in rat TS cells cultured in undifferentiated states. However, Mmp9 and Mmp10 increased expression in TS cells following exposure to deferoxamine (Fig. 6a; P < 0.05), which is similar to the upregulation of these genes in placentas exposed to iron deficiency. Therefore, deferoxamine treatment of rat TS cells is sufficient to recapitulate some aspects of placental gene expression changes associated with maternal iron deficiency in vivo.
Figure 6.
Increased expression of junctional zone markers in rat TS cells cultured with an iron chelator, deferoxamine. Rat TS cells were incubated in the presence or absence of 25μM or 50μM deferoxamine (DFO) for 48 hours. (a) qRT-PCR analysis showing transcript levels of Cdx2, Rbp4, Mmp9, Mmp10, Prl3d1, and Tpbpa in rat TS cells cultured with and without DFO. (b) Immunofluorescence was used to detect TPBPA in rat TS cells exposed to 0, 25, or 50μM DFO. DAPI was used to stain nuclei. Results from qRT-PCR are presented as means ± SEM, with P values reflecting 1-way ANOVA with Tukey’s post hoc test. An asterisk (*) denotes statistical significance (P < 0.05, n = 3-4). Scale bar = 50 μm.
Since an expanded junctional zone was a consistent feature of placentas exposed to maternal iron deficiency, we investigated whether exposure of rat TS cells to deferoxamine would stimulate expression of genes associated with junctional zone development. Interestingly, the junctional zone marker Trophoblast-Specific Protein Alpha (Tpbpa) and trophoblast giant cell marker Prolactin Family 3, Subfamily d, Member 1 (Prl3d1 – encodes Placental Lactogen-I) were dose-dependently increased in the presence of deferoxamine, whereas there was no change in expression of the TS cell stem state marker, Caudal-type Homeobox-2 (Cdx2,Fig. 6a). Increased TPBPA was also evident in rat TS cells exposed to deferoxamine at the protein level, as determined by immunofluorescence (Fig. 6b). These findings indicate that TS cell function is altered in conditions of reduced iron levels, preferentially driving development of junctional zone lineages.
Discussion
In this study, we used a rat model to investigate the effects of maternal iron deficiency on placental structure and development. To summarize, we report that maternal iron deficiency and anemia resulted in: (1) asymmetrical fetal growth restriction as demonstrated by decreased fetal body weight and liver weight but no change in brain, heart, or kidney weight; (2) increased placental weight and junctional zone area near the end of gestation; (3) altered expression of placental genes including those associated with nutrient and oxygen transport, lipoprotein metabolism, hormone response, and cell communication; and (4) robust changes in levels of proteins associated with iron storage and homeostasis despite mild effects on expression of genes that encode these proteins. We also report that TS cells cultured in conditions of low iron availability are primed to differentiate into junctional zone lineages. Taken together, these findings suggest that maternal iron deficiency results in fetal growth restriction, placental hypertrophy, and altered expression of placental genes, which may have important implications in the programming of long-term health in offspring (5).
The rat model used in this study is one in which iron deficiency develops in the dam before gestation and progresses to show signs of anemia during pregnancy. This model is intended to mimic a common clinical scenario in which the increased iron demands of pregnancy coupled with insufficient iron intake or poor gastrointestinal absorption causes gradual depletion of iron stores, resulting in clinical manifestations of anemia near the end of gestation (25). Our model recapitulates the clinical scenario well, as maternal Hb levels before gestation reside within the range considered normal in nonpregnant women (12-16 g/dL), and results in anemia as pregnancy progresses (<11 g/dL) (44).
Maternal iron restriction caused changes in fetal iron status and growth trajectories. Maternal iron deficiency was associated with decreased fetal Hb levels in late gestation, suggesting that there is inadequate supply of iron available for the fetus by the end of pregnancy. Interestingly, there was no change in relative levels of ferritin in GD18.5 fetal livers from dams fed iron-replete and iron-deficient diets. This was unexpected given the close association between serum ferritin levels and iron availability but could indicate a compensatory mechanism by the fetus to sequester iron in conditions of poor maternal iron availability. Alternatively, ferritin is induced during inflammation and low oxygen availability (45, 46). Given the poor placental development and reduced fetal growth evident in this model, liver ferritin levels may not be a reliable indicator of iron availability in fetuses exposed to maternal iron deficiency. Iron-deficient fetuses also exhibited growth restriction as indicated by decreased total body weight and reduced relative liver weight but not brain weight, which is consistent with findings reported in other rodent models of maternal iron deficiency (22-26). As iron-deficient dams exhibited no change in cumulative food intake and body weight gain, fetal growth effects can most likely be attributed to a lack of iron and not altered intake of calories or other micronutrients in the maternal diet. Decreased fetal growth may have important implications for short-term survival and long-term cardiometabolic health in offspring (47).
Placental modifications were apparent in iron-deficient dams during late gestation. Placentas from iron-deficient dams on GD18.5 displayed increased absolute and relative weights, consistent with findings from other rodent models of iron deficiency (23-25, 27, 28) and human cases of anemia (11, 13-17). When compared to fetal weight, placental weight can also be used as a proxy for placental efficiency, and an indication of how placental development and function has adapted to sustain fetal requirements (48). Whether such placental modifications are adaptive responses striving to compensate for iron deficiency and/or impaired oxygen delivery, or pathological changes contributing to fetal adversity remains an area of further investigation.
Although a change in absolute or relative placental size does not necessarily indicate altered placental function, additional insight can be provided by determining whether elements of placental structure are altered. No significant change was detected in labyrinth zone area between iron-deficient and control pregnancies. However, placentas from iron-deficient dams exhibited increased junctional zone area. Comprising trophoblast giant cells, spongiotrophoblast cells, glycogen cells, and invasive trophoblast cells, the junctional zone forms the interface between the placenta and decidua basalis, constitutes the main endocrine compartment of the rodent placenta, and promotes nutrient flow to the conceptus. An expanded junctional zone area may indicate a compensatory response to decreased nutrient (iron) or gas (oxygen; due to reduced Hb) availability in iron deficiency. In vivo and in vitro experiments have previously demonstrated that allocation of trophoblasts into junctional zone lineages is induced by low oxygen conditions, which may explain why increased junctional zone area was evident in anemic rats (49, 50). Indeed, gene ontology analysis identified oxygen transport as a biological pathway upregulated in placentas during maternal iron deficiency. Moreover, we found that the iron chelator deferoxamine (which can stimulate cellular hypoxic responses) primed rat TS cells to form junctional zone lineages as shown by increased expression of Tpbpa and Prl3d1, suggesting that the expanded junctional zone in maternal iron deficiency may be a compensatory response by trophoblasts to accelerate oxygen and nutrient flow to the placenta. Since human extravillous trophoblasts perform analogous functions as rodent junctional zone trophoblasts in terms of promoting oxygen and nutrient delivery to the placenta, it is tempting to speculate that extravillous trophoblasts may respond similarly in human cases of maternal iron deficiency, especially given that their development is facilitated during culture in low oxygen (51).
Analysis of both GD13.5 and GD18.5 provided insight into the timing of placental modifications. Interestingly, changes in placental weight and morphology between iron-deficient and control pregnancies were only detected on GD18.5, indicating placental structural changes do not become evident until later in gestation. This was unexpected, given that the zonal architecture of the placenta is established by GD13.5. Although we did not observe changes in placental weight or structure on GD13.5, it is possible that maternal iron deficiency causes more subtle changes in placentation during early or mid-gestation. Nevertheless, since placental structure was obviously affected by iron deficiency on GD18.5, we investigated gene expression changes in GD18.5 placentas. Numerous differentially expressed genes were apparent in placentas exposed to maternal iron deficiency compared to controls. Those with the greatest fold increase included Alox15 and Mmp10. ALOX15 is an enzyme involved in lipid oxygenation, generating products that regulate inflammation and immunity (52-55). ALOX15 has anti-inflammatory effects including reduction of leukocyte migration, apoptosis of proinflammatory neutrophils and differentiation of M2 macrophages (56-59). Therefore, increased expression of Alox15 may be a response to dampen inflammation in placentas exposed to maternal iron deficiency (60). Increased ALOX15 expression has also been detected in other tissues during iron deficiency, including liver, intestine, and pancreas, coinciding with altered profiles of biologically active lipid mediators in these tissues (61, 62). Whether placentas subjected to iron deficiency also possess altered profiles of lipid mediators due to increased ALOX15 expression is not yet known and is the subject of our future investigation. However, maternal iron deficiency in rats is associated with altered lipid profiles in maternal plasma as well as maternal and fetal liver, supporting the notion that iron deficiency has profound impacts on lipid metabolism and fetal programming (63). Interestingly, in pregnant rats exposed to streptozotocin-induced diabetes, Alox15 mRNA is induced in the junctional zone, suggesting that a variety of nutrient stresses may upregulate ALOX15 in this tissue (64). MMP10 is a member of a metalloprotease family highly linked with cell invasion and tissue remodeling, including extravillous trophoblasts (65-67). Upregulation of Mmp10 may lead to increased matrix remodeling in iron-deficient placentas, facilitating trophoblast invasion to ensure the growing conceptus receives a sufficient supply of maternal blood. This finding is further supported by our gene ontology and pathway analysis, which detected a change in cell adhesion and cell communication pathways in placentas from iron-deficient dams.
Genes with the greatest fold decrease included Prl3a1 and Apoa1. Prl3a1 encodes a placental prolactin-related hormone expressed by spongiotrophoblast cells in the junctional zone, with implications in fetal growth and development (68). Decreased Prl3a1 expression in GD18.5 mouse placentas has previously been associated with fetal growth restriction resulting from maternal protein restriction (69). Apoa1 encodes an apolipoprotein involved in lipid transport. Decreased expression of Apoa1 may signify impaired lipid homeostasis (70, 71), and is consistent with the concept that iron deficiency impairs maternal and fetal lipid metabolism and restricts the supply of fatty acids to the fetus (23, 63, 72). These results and their implications for fetal growth are further supported by our gene ontology and pathway analysis, which detected downregulation of lipoprotein metabolism and vitamin transport.
In summary, our findings demonstrate that maternal iron deficiency and anemia results in fetal growth restriction, placentomegaly, expanded junctional zone size, and altered expression of genes within the placenta, which may have important implications in placental function, fetal growth, and long-term health of offspring. Future studies are needed to clarify molecular mechanisms in the placenta during maternal iron deficiency and determine whether these modifications are adaptive responses striving to maintain iron sustenance and oxygen delivery in order to support fetal growth and development, or pathological changes contributing to maternal and fetal adversity. Moreover, there may be important differences in placental responses that were not captured in our analysis, such as the severity of anemia, timing of onset, and fetal/placental sex, all of which may influence placental adaptability or pathophysiology. For example, mouse placental responses to iron deficiency show robust sexual dimorphism, and it would be of interest to determine whether the junctional zone changes observed in our study differ depending on whether the placenta is male or female (38). Understanding how iron deficiency impacts placental and fetal health may help to uncover earlier interventions and improved fetal outcomes.
Acknowledgments
The authors wish to thank Michael Soares (University of Kansas Medical Centre, Kansas City, KS) for providing the rat TS cells.
Financial Support: This study was funded through grants awarded from the Canadian Institutes of Health Research (MOP142396 to S.L.B.; PJT376512 to S.J.R.) and the Ontario Early Researcher Awards (to S.J.R.) A.G.W. was supported by a Vanier Canada Graduate Scholarship from the Canadian Institutes of Health Research and an Alberta Innovates Graduate Scholarship; M.J.J. was supported by an Alexander Graham Bell Canada Graduate Doctoral Scholarship from the Natural Sciences and Engineering Research Council of Canada.
Author Contributions: H.R., A.G.W., S.L.B., and S.J.R. contributed to the overall approach and design of experiments. A.G.W. conducted all experiments involving the use of animals, tissue collection and weighing, and Hb assessments. H.R., K.J.B., and M.J.J. performed all other experiments and performed data analysis. H.R., S.L.B., and S.J.R. wrote the manuscript. All authors critically revised the manuscript and approved the final version.
Glossary
Abbreviations
- ANOVA
analysis of variance
- DAPI
4′,6-diamidino-2-phenylindole
- GD
gestational day
- Hb
hemoglobin
- PBS
phosphate-buffered saline
- qRT-PCR
quantitative reverse-transcriptase polymerase chain reaction
- SDS
sodium dodecyl sulfate
- SEM
standard error of the mean
- TPBPA
trophoblast specific protein alpha
- TS
trophoblast stem
Additional Information
Disclosures: The authors have nothing to disclose.
Data Availability
Some or all data generated or analyzed during this study are included in this published article or in the data repositories listed in References. The Clariom S dataset is available at the National Center for Biotechnology Information Gene Expression Omnibus under accession GSE163226.
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Associated Data
This section collects any data citations, data availability statements, or supplementary materials included in this article.
Data Availability Statement
Some or all data generated or analyzed during this study are included in this published article or in the data repositories listed in References. The Clariom S dataset is available at the National Center for Biotechnology Information Gene Expression Omnibus under accession GSE163226.