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 Table of Contents  
ORIGINAL ARTICLE
Year : 2020  |  Volume : 4  |  Issue : 3  |  Page : 137-145

Differentiation of human adipose-derived stem cells into endometrial epithelial cells


Department of Obstetrics and Gynecology, Reproductive Medical Center, Tangdu Hospital, The Fourth Military Medical University, Xi'an 710038, China

Date of Submission01-Jun-2020
Date of Decision13-Jan-2020
Date of Acceptance10-Aug-2020
Date of Web Publication29-Sep-2020

Correspondence Address:
Xi-Feng Xiao
Department of Obstetrics and Gynecology, Reproductive Medical Center, Tangdu Hospital, The Fourth Military Medical University, 569 Xinsi Road, Baqiao District, Xi'an 710038
China
Xiao-Hong Wang
Department of Obstetrics and Gynecology, Reproductive Medical Center, Tangdu Hospital, The Fourth Military Medical University, 569 Xinsi Road, Baqiao District, Xi'an 710038
China
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Source of Support: None, Conflict of Interest: None


DOI: 10.4103/2096-2924.296547

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  Abstract 


Objective: This study aimed to investigate the differentiation of human adipose-derived stem cells (hASCs) into endometrial epithelial cells (EECs) under certain induction conditions and to a further step provide a promising approach for ASCs in clinical practice to the treatment of severe intrauterine adhesion.
Methods: Four groups of hASCs were separately cultured as follows: in Group 1, hASCs were cultured in a control medium (5% fetal bovine serum [FBS] + α-minimum Eagle's medium [α-MEM]); in Group 2, hASCs were cultured in an induction medium (5% FBS + α-MEM + [1 × 10−7 mol/L 17β-estradiol] + 10 ng/mL transforming growth factor β1 [TGF-β1] + 10 ng/mL epidermal growth factor [EGF] + 10 ng/mL platelet-derived growth factor BB [PDGF-BB]); in Group 3, hASCs and human endometrium cells (hEMCs) were cocultured in the control medium; and in Group 4, hASCs and hEMCs were cocultured in the induction medium.
Results: When cocultured with hEMCs, the morphology of hASCs became similar with EECs, and the addition of factors such as EGF, TGFβ, PDGF-BB, and 17β-estradiol promoted differentiation. This study, for the first time, demonstrated estrogen receptor (ER)α and ERβ expression in hASCs and preliminarily explored changes in ERα, ERβ, β-catenin, and H19 mRNA expression during hASC differentiation. Furthermore, we concluded that H19 mRNA expression was negatively correlated with differentiation, which is seemingly related to the estrogen signaling pathway.
Conclusions: hASCs revealed the potential for differentiating to EECs when cocultured with hEMCs.

Keywords: Adipose-derived Stem Cell; Endometrial Epithelial Cell; Estrogen Receptor α; Estrogen Receptor β; H19; β-catenin


How to cite this article:
Yang F, Zhang WL, Chen SQ, Sun HJ, Lu J, Xiao XF, Wang XH. Differentiation of human adipose-derived stem cells into endometrial epithelial cells. Reprod Dev Med 2020;4:137-45

How to cite this URL:
Yang F, Zhang WL, Chen SQ, Sun HJ, Lu J, Xiao XF, Wang XH. Differentiation of human adipose-derived stem cells into endometrial epithelial cells. Reprod Dev Med [serial online] 2020 [cited 2020 Oct 22];4:137-45. Available from: https://www.repdevmed.org/text.asp?2020/4/3/137/296547




  Introduction Top


Intrauterine adhesions (IUAs) mostly occur following the curettage of the uterus during early pregnancy.[1] Other etiologies include myomectomy, hysteroscopic resection of uterine septum/conception remnants, radiation therapy for gynecological cancer, uterine artery embolization for postpartum hemorrhage, genital tuberculosis infection, and cesarean section.[2] IUAs present with a wide spectrum of clinical manifestations corresponding to the extent of scarring inside the uterine cavity. Some of the clinical conditions associated with IUAs include menstrual fluctuations such as amenorrhea and hypomenorrhea, infertility, placental abnormalities, and chronic pelvic pain. Hysteroscopic adhesiolysis is the primary treatment aimed at restoring the shape of the uterine lining. Unfortunately, surgical treatment for severe IUAs is complicated by a high rate of recurrent adhesions, although often assisted with several adjuvant therapies, such as physical barriers, postoperative estradiol supplementation, and second-look hysteroscopy.[3]

In recent years, stem cell-based bioengineering strategies have offered new hope to overcome the treatment bottleneck for IUAs. Stem cells are undifferentiated cells that can self-renovate and differentiate into multiple lineages. Mesenchymal stem cells (MSCs) isolated from adult tissues are immunocompatible and are not subject to ethical challenges;[4] therefore, these are the most suited for clinical applications. This novel approach has demonstrated enormous potential to treat massive IUAs as well as stubborn thin endometrium responsible for infertility and repeated implantation failure inin vitro fertilization and embryo transfer.[5] MSCs derived from the bone marrow, endometrium, menstrual blood, umbilical blood, amnion, and adipose tissue contribute to endometrial regeneration.[6],[7],[8],[9],[10],[11],[12] Among these MSCs, adipose-derived stem cells (ASCs) have drawn much attention owing to their easy accessibility, high cell yields, highin vitro proliferation, weak immunogenic effect, and high multilineage differentiation capacity.[13],[14] Ye et al. and our group have previously demonstrated the therapeutic effects of ASCs on endometrial regeneration in a rat model.[10],[15]

Nonetheless, the in-depth mechanisms of ASC repair in endometrial damage remain debatable and warrant further research. Endometrial repair depends on regenerative mechanisms that lead to the reconstruction of damaged functional endometrial layer. Cell fusion, paracrine effects, immunomodulatory effects, microvesicles carrying mRNAs or microRNAs, and mitochondrial transfer are some of the recent candidates in discussion regarding the curative effect of MSCs.[16],[17],[18] Moreover, ASCs may serve as a source of antioxidants, free radical scavengers, and chaperone/heat shock proteins and also as the stem cell niche of the host by stimulating the recruitment of endogenous stem cells to the damaged areas and their commitment in the proper lineage.[13],[19] However, the major value of ASCs is primarily related to their multilineage differentiation potential. ASCs support adipogenesis,[20] osteogenesis,[21] chondrogenesis,[22] and myogenesis[23] under specific induction conditions. Other cell lineages differentiated from ASCs include neuron-like cells,[24] hepatocytes,[25] endothelial-like cells,[26] and islet-like cells.[27] Furthermore, ASCs can differentiate into other epithelial lineages;[28],[29] however, evidence regarding the differentiation potential of ASCs into endometrial epithelial cells (EECs) is limited.

In this study, we hypothesized that human ASCs (hASCs) can differentiate into EECs when cocultured with mature human endometrium cells (hEMCs) in a differentiation medium (transforming growth factor β1 [TGF-β1], epidermal growth factor [EGF], platelet-derived growth factor BB [PDGF-BB], and 17β-estradiol). Moreover, we preliminarily investigated the molecular pathways involved in differentiation. To the best of our knowledge, this study provides the first evidence for the expression of estrogen receptors α and β (ER-α and ER-β, respectively) in hASCs. Further insight into these mechanisms would help to provide a promising approach for ASCs in clinical practice to the treatment of severe IUA.


  Methods Top


Patient information

The research was approved by the Institutional Review Board of Tangdu Hospital, Fourth Military Medical University (approval number: 201712-01). We isolated hASCs from the abdominal subcutaneous adipose tissues, which were discarded during general surgeries, of reproductive-age women, as described previously.[30] The discarded abdominal subcutaneous adipose tissues were from 13 patients with average age of 30.1 years old, among whom 4 were undergoing surgery for ovarian endometriosis cyst, 4 were undergoing surgery for ectopic pregnancy, and 5 were undergoing surgery for benign ovarian tumor. hEMCs were isolated from biopsied endometrial tissue of 9 women with an average age of 31.3 years old, among whom 5 were primary infertility and 4 were secondary infertility.

Isolation and culture of human adipose-derived stem cells and hEMCs

The adipose tissue was thoroughly washed in phosphate-buffered saline (PBS) (Corning, USA) and was sliced into small clumps (~1.0 mm3). Then, these clumps were enzymatically digested for 90 min at 37°C in a 5% CO2 incubator using collagenase Type I (Sigma, USA) at a working concentration of 0.2 mg/mL (Thermo, USA). The digested adipose tissue was gently blew evenly using a pipette. To stop the digestion, an adequate amount of complete α-minimum Eagle's medium (α-MEM) was added (Gibco, USA) (10 g/L α-MEM powder, 2 g/L sodium bicarbonate, and 10% fetal bovine serum [FBS] [Sijiqing, China], 100 U/mL penicillin, and 100 μg/mL streptomycin). The suspension was successively filtered through 150- and 75-μm metal meshes to remove cellular debris. Then, the filtrate was centrifuged at 800 ×g for 5 min, and the mature adipose-containing supernatant was discarded. The precipitate was identified as the stromal vascular fragment (SVF) and resuspended in 3 mL α-MEM culture medium. This suspension was centrifuged again at 800 ×g for 5 min. The supernatant was discarded, and portions of SVF were resuspended in 5 mL α-MEM cell culture medium. Then, cells were seeded in a 25-cm2 flask and cultured at 37°C in 5% CO2. The medium was first changed after an initial 24 h of culture and then changed every other day until the adherent cells were split upon achieving 80% confluence for passage. This initial passage of the primary cell culture was referred to as passage 0 (P0). Cells were passaged by trypsin (0.25%, 3–5 min) digestion and seeded in flasks at 1:3 ratio repeatedly after achieving a density of 75%–90%. P3–P8 hASCs were used for all experiments. All cell culture steps were strictly in accordance with the aseptic principle.

hEMCswere isolated from biopsied endometrial tissue of reproductive-age women undergoing infertility examination at 4–5 days of the menstrual cycle. The steps involved tissue cutting, digestion, suspension, centrifugation, and cell seeding, similar to those for hASCs. However, the duration of tissue digestion immediately after cutting was extended to 120 min, and the speed and duration of supernatant centrifugation were increased to 1000 ×g and 10 min, respectively. Cells were seeded onto 6-well plates (Thermo, USA). The subsequent coculture with hASCs utilized the P0 cells.

Hematoxylin and eosin staining

The standard protocol of hematoxylin and eosin (H and E) staining was followed to examine the morphology of EECs. Briefly, the culture medium residue was rinsed thrice in PBS. Then, cell slides were dehydrated using xylene, absolute alcohol, and 75% alcohol in that order and then soaked in hematoxylin for 5 min. Before staining with eosin, the cell slides were washed and dehydrated in gradient alcohol (85% and 95% for 5 min, respectively). To make the cell sample transparent after eosin staining, absolute alcohol and xylene were successively used. Furthermore, neutral resin was used to seal the slides for further observation under an inverted microscope (ZEISS, Germany).

Cellular proliferation assay

hASC proliferation was assessed using the cell counting kit-8 (CCK8) assay (Peprotech, USA). The assay was performed according to the manufacturer's instructions. Briefly, P3 cells were seeded (2 × 103 cells/well) into seven 96-well plates and cultured at 37°C in 5% CO2. Every 24 h, 10 μL of CCK-8 working solution (50 μg/mL Vc) was added to each well and incubated for 1–4 h. To eliminate the effect of Vc on the CCK-8 solution, controls were set wherein the culture medium or CCK-8 and Vc were added to wells without cells. Absorbance at 450 nm was measured with a microplate reader (MD, USA). All experiments were performed in triplicate.

Flow cytometry

Flow cytometry was used to phenotypically evaluate hASCs for the expression of CD29, CD44, CD90, CD105, and CD45. P3 hASCs in the logarithmic growth period were harvested. The precipitate was resuspended with a wash solution (5% FBS, 2 g/L sodium azide, and PBS). The resulting cell suspension (5 × 105 cells/L) was aliquoted into six tubes (Millipore, USA) for centrifugation (1 mL/tube). The supernatant was discarded, and 4 μL each of monoclonal (CD29-PE, CD44-APC, CD45-percp-cy5.5, CD90-FITC, and CD105-APC [Peprotech, USA]) and control antibodies were added into each previously labeled tube and incubated for 30 min at 4°C in the dark. Then, the unbound antibodies washed off, and cells were fixed using 400 μL of fixing liquid (0.2 g/L sodium azide, 20 g/L glucose, and 1% paraformaldehyde [PFA] [Ke Hao Bioengineering Co., Ltd., China]) and PBS. Debris was filtered through a 75-μm metal mesh before analysis. Thereafter, the prepared samples were tested for target cell surface markers using flow cytometry (BD, UDA). Data were analyzed using FlowJo 7.6 (LLC Ashland, OR, USA).

Adipogenic and osteogenic induction and identification

P3 hASCs in the logarithmic growth period were harvested and seeded into 6-well plates (2 × 104 cells/well). After 24 h of culture in the α-MEM medium when the cells became adherent, the culture medium was changed to an adipogenic-inducing medium (10% FBS, 1% streptomycin, 1% penicillin, 1 mol/L dexamethasone, 0.1 mmol/L indomethacin, 10 mg/L insulin, 0.5 mmol/L 3-isostationary-1-methyl xanthine, and α-MEM medium). The inducing medium was replaced every other day until the cell culture lasted for 7 days. The success of adipogenic induction was assessed by oil red O staining. Then, the culture medium residue was washed twice with PBS. Oil red O was then added into each well and stained for 60 min, and the dye was washed off with PBS for subsequent observation under an inverted microscope.

To induce osteogenic differentiation, cells were exposed to an osteogenic-inducing medium (10 mol/L dexamethasone, 50 μg/mL vitamin C, 10 mmol/L β-sodium glycerophosphate, 10% FBS, 1% streptomycin (Thermo, USA), 1% penicillin (Thermo, USA), and α-MEM medium). The inducing medium was replaced every 48 h until the cell culture lasted for 3 weeks. The success of osteogenic induction was confirmed by Alizarin red staining. Culture medium residue was routinely washed off, and Alizarin red was added to each well and stained for 10 min. Then, the dye was washed off by PBS for subsequent observation under an inverted microscope.

Differentiation of human adipose-derived stem cells toward endometrial epithelium

To induce differentiation of hASCs toward endometrial epithelium, the transwell system (0.4 μm, Corning, USA) was employed. P0 hEMCs in logarithmic growth period (3,000 cells) were harvested and seeded as the top layer membrane of the transwell chamber. The culture medium was changed the next day to remove the cell debris in the supernatant, and P3 hASCs were then seeded on the cover slips placed at the bottom of 6-well plates. After 4 h, when the hASCs became adherent, the top layer of the transwell with hEMCs was moved to each well of the 6-well plate. Then, hASCs and hEMCs were cocultured for 5 days.

To evaluate the outcome of induction, the following four groups were set: in Group 1, hASCs were cultured in the control medium (5% FBS + α-MEM); in Group 2, hASCs were cultured in the induction medium (5% FBS + α-MEM + [1 × 10−7 mol/L 17β-estradiol] [Sigma, USA] + 10 ng/mL TGF-β1 [Peprotech, USA] + 10 ng/mL EGF [Peprotech, USA] + 10 ng/mL PDGF-BB [Peprotech, USA]); in Group 3, hASCs and hEMCs were cocultured in the control medium; and in Group 4, hASCs and hEMCs were cocultured in the induction medium. The concentration of 17β-estradiol involved in the induction medium was referred to the previous articles and pre experiment. Shi et al.[11] used 17β-estradiol at 1 × 10−6 mol/L 17β-estradiol in their experiment assessing the differentiation of human umbilical cord Wharton's jelly-derived MSCs into endometrial cells. Zhang et al.[31] showed that CK7 and CK19 expression levels were highest in the 1 × 10−8 mol/L 17β-estradiol group in thein vitro study on differentiation of bone marrow MSCs into EECs in mice. Our pre experiment used three 17β-estradiol concentrations, namely, 1 × 10−8 mol/L, 1 × 10−7 mol/L and 1 × 10−6 mol/L, and the result showed that the concentration at 1 × 10−7 mol/L resulted in the highest expression of CK18. Therefore, we used 1 × 10−7 mol/L as the final 17β-estradiol concentration in the induction medium.

Immunofluorescence staining

Immunofluorescence (IF) staining was used to detect the expression of target markers in endometrial cells (CK18, vimentin, and ER-α), hASCs (ER-α and ER-β), and differentiated hASCs (CK18). The cells cultured in 6-well plates or cover slips were first fixed in 4% PFA for 15 min at room temperature and then washed three times with PBS. The cells were then permeabilized with 0.5% Triton X-100 (dissolved in PBS; Sigma, USA) for 20 min and incubated with 3% hydrogen peroxide for 10 min. Then, these cells were washed three times for 3 min with PBS, and the PBS residue was soaked up by an absorbent paper. Thereafter, the cells were blocked with ready-to-use normal goat serum (Bolster, China) for 30 min at room temperature. Next, the cells were incubated with the following primary antibodies in a wet box at 4°C overnight: CK18 (rabbit, 1:500; Abcam), vimentin (rabbit, 1:1,000; Abcam), ER-α (human, 1:200; Peprotech, USA), ER-β (human, 1:200; Peprotech, USA), and progesterone receptor (PR) (human, 1:1,000; Peprotech, USA). Next, the plates were washed three times for 5 min with PBS and soaked up before adding the secondary antibodies. The secondary antibodies were the corresponding fluorescence-labeled immunoglobulin G with the primary antibodies (all from Peprotech, USA). Cell were incubated with secondary antibodies in a wet box for 1 h and then washed and soaked as in the previous step. Subsequently, DAPI (Sigma, USA) was added, and cells were incubated for 5 min to stain the cell nuclei. Excess dye was washed with PBS and soaked up as before. Finally, the prepared slips were sealed with a fluorescent mounting medium and observed under an inverted fluorescent microscope.

Quantitative real-time polymerase chain reaction analysis

Quantitative real-time polymerase chain reaction (qRT-PCR) was performed to test the mRNA expression changes of the concerned molecules during the differentiation of hASCs to EECs. Total RNA was isolated from P3 hASCs at the beginning and end of culture induction using Trizol® (SIGMA, USA) and then reverse-transcribed into cDNA using the RT kit (Applied Biological Materials, Italy). QuantiNova™ SYBR® Green PCR Kit (QIAGEN, Germany) was used for qRT-PCR following the manufacturer's instructions. PCR was run on Bio-Rad PCR machine (USA) using a reaction mixture containing 10 μL of 2× SYBR Green PCR Master, 0.75 μL (0.1 μmol/L)of primer A+, 0.75 μL (0.1 μmol/L) of primer B+, 7 μL of RNase-free water (Solarbio, China), and 1.5 μL of template cDNA under the following conditions: denaturation at 95°C for 2 min, annealing at 95°C for 5 s, and extension at 60°C for 10 s; reactions were repeated for 40 cycles. The target gene expression levels were determined using the 2ΔΔCt method for the relative quantification of gene expression. All experiments were performed in triplicate.

All primers were synthesized by Sangon Biotech (China): ER-α forward: 5'-CCCACTCAACAGCGTGTCTC-3'; ER-α reverse:

5'-CGTCGATTATCTGAATTTGGCCT-3'; ER-β forward:

5'-AGCACGGCTCCATATACATACC-3'; ER-β reverse:

5'-TGGACCACTAAAGGAGAAAGGT-3'; H19 forward:

5'-GGCAAGAAGCGGGTCTGT-3'; H19 reverse:

5'-GTGCAGCATATTCATTTCCAAG-3'; β-catenin forward:

5'-AAAGCGGCTGTTAGTCACTGG-3'; β-catenin reverse:

5'-GTGCAGCATATTCATTTCCAAG-3'; β-actin forward:

5'-AGCGAGCATCCCCCAAAGTT-3'; and β-actin reverse:

5'-GGGCACGAAGGCTCATCATT-3'.

Statistical analysis

Statistical analysis was performed using SPSS19.0 (IBM®, USA). Quantitative data were presented as mean ± standard deviation. Differences between groups were analyzed by independent sample t- test, and P < 0.05 was considered statistically significant.


  Results Top


Growth and identification of human adipose-derived stem cells

After 4 h of culture, P0 hASCs started to become adherent, and after 24 h of culture, the cells became completely adherent. Cell proliferation was extremely slow, and a small short shuttle or polygon shape was observed in the first 4 days of culture [Figure 1]a. The cell volume significantly increased at day 5, and the cells rapidly cloned after 6 days of culture. Adherent cells were split upon achieving 90% confluence for passage on days 8–9. The cell size and shape tended to be more consistent with subsequent passages. After P3, the cells seemingly appeared as fibroblasts and grew rapidly to achieve a density of 75%–90% only in 6–7 days [Figure 1]b. The growth pattern of P3 hASCs was as follows: on days 1–2 of cell seeding, the cells slowly proliferated during the latent period; on days 3–5, cell growth entered a rapid logarithmic growth period; and on days 6–7, the cell growth reached a plateau; the recessionary period followed thereafter [Figure 1]c.
Figure 1: Identification of hASC through morphology of hASCs observed under an inverted microscope (×50) (a and b), growth pattern of P3 hASCs (c), flow cytometry (d), adipogenesis (e) and osteogenesis (f). Sample number = 13. hASC: Human adipose-derived stem cells.

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Regarding the identification of hASCs, flow cytometry showed that in hASCs, the expression of CD29, CD44, CD90, and CD105 was positive, whereas that of the hematopoietic stem cell marker CD45 was negative [Figure 1]d. Adipose droplets formed within hASCs after adipogenic induction and appeared red when stained with Oil red O [Figure 1]e. Reddish-brown mineralized nodules were also observed on the surface of hASCs through osteogenic induction culture when stained with Alizarin red [Figure 1]f.

Morphology and expression of hEMC molecular markers

After 2 h of culture, P0 hEMCs started to become adherent, and after 24 h of incubation, nearly 80% of the hEMCs became adherent. The supernatant containing lymphocytes and debris was discarded. Human endometrial cells are a group of heterogenously adherent cells, mainly including EECs and endometrial stromal cells (ESCs). The morphologies of EECs and ESCs were distinct. On day 3 of culture, the shape of P0 EECs resembled that of tadpoles, and the cells grew in a whirl manner with a clear boundary [Figure 2]a. Meanwhile, ESCs were spindle-shaped or polygonal and were scattered evenly without showing any trend in direction [Figure 2]b. As confirmed by H and E staining, both cell types possessed abundant cytoplasm and an elliptical nucleus [Figure 2]c.
Figure 2: Morphology of EECs and ESCs observed under an inverted microscope. After 3 days of culture, the shape of P0 EECs resembled that of tadpoles, and they grew in a whirl manner with a clear boundary (×100) (a), whereas the ESCs were spindle- or polygon-shaped and were scattered evenly without showing any trend in direction (×100) (b). H and E staining showed that both cell types possessed abundant cytoplasm and an elliptical nucleus; black arrows indicate EECs (c). IF staining showed that CK18 was specifically expressed in EECs (d). Vimentin was simultaneously expressed in EECs and ESCs (e). ERs (f) and PRs (g) were positively expressed in both cell types, but the expression of PR was considerably weaker (g) than that of ER-α. Sample number = 9. Differences between groups were analyzed by independent sample t-test. EECs: Endometrial epithelial cells; ESCs: Endometrial stromal cells; H and E: Hematoxylin and eosin; IF: Immunofluorescence; ERs: Estrogen receptors; PRs: Prgestrone receptors.

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As revealed by IF staining, CK18 was specifically expressed in EECs [Figure 2]d, whereas vimentin was simultaneously expressed in both EECs and ESCs [Figure 2]e. While ERs [Figure 2]f and PRs [Figure 2]g were positively expressed in both cell types, the expression of PR was considerably weaker than that of ER-α [Figure 2]g.

Outcome of the differentiation of human adipose-derived stem cells into endometrial epithelial cells

The expression of epithelium lineage-specific marker CK18 was examined by IF staining of hASCs in four treatment group (see Methods section) following differentiation. CK18 was negatively expressed in Group 1 and 2 hASCs, which were cultured alone. Interestingly, CK18 was positively expressed in Group 3 and 4 hASCs, which were cocultured with hEMCs. Moreover, Group 4 hASCs were cocultured with hEMCs on the induction medium; the morphology of these cells was the most similar to that of mature human EECs [Figure 3]a.
Figure 3: IF (a) showed that CK18 was negatively expressed in Group 1 and 2 hASCs but positively expressed in Group 3 and 4 hASCs. The morphology of hASCs in Group 4 was the most similar to that of mature human EECs. qRT-PCR results (b) showed that the level of CK18 mRNA expression consecutively increased in the four groups with statistically significant differences. Meanwhile, the mRNA expression levels of CK7, CK19, and EMA also increased in Groups 2, 3, and 4, but without statistically significant differences. Sample number = 13. Differences between groups were analyzed by independent sample t-test. hASC: Human adipose-derived stem cells; qRT-PCR: Quantitative real-time polymerase chain reaction; IF: Immunofluorescence.

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Furthermore, the mRNA expression levels of CK7, CK18, CK19, and EMA were measured via qRT-PCR. The level of CK18 mRNA expression consecutively increased in the four groups with statistically significant differences. Similarly, the level of CK7, CK19, and EMA mRNA expression also increased in Groups 2, 3, and 4, although without statistically significant differences [Figure 3]b.

Changes in ER-α, ER-β, β-catenin, and H19 mRNA expression levels during differentiation

As shown in IF staining, ER-α and ER-βwereinitially expressed both in the nucleus and cytoplasm of hASCs, but mainly in the cytoplasm [Figure 4]a. qRT-PCR results showed that the levels of ER-β and β-cateninmRNA expression steadily increased from Group 1 through Group 4 with statistically significant differences. Although the expression of ER-α increased from Group 1 to Group 4, the differences were not statistically significant. Interestingly, the level of H19 mRNA expression was lower in Group 2 than in Group 1. A similar trend was observed between Groups 3 and 4; however, in the presence of the induction medium containing growth factors and 17β-estradiol, the expression of H19 mRNA was significantly downregulated [Figure 4]b.
Figure 4: IF (a) showed that ER-α and ER-β were initially expressed both in the nucleus and cytoplasm of hASCs, but mainly in the cytoplasm. qRT-PCR results (b) showed that the mRNA expression levels of ER-β and β-catenin steadily increased from Group 1 through Group 4, with statistically significant difference. The mRNA expression levels of ER-a also increased from Group 1 to Group 4. However, the expression level of H19 mRNA was lower in Group 2 than in Group 1 and lower in Group 4 than in Group 3, suggesting that in the presence of induction medium containing growth factors and 17β-estradiol, the expression of H19 mRNA was significantly downregulated. Sample number = 13. Differences between groups were analyzed by independent sample t-test. IF: Immunofluorescence; hASC: Human adipose-derived stem cells; qRT-PCR: Quantitative real-time polymerase chain reaction; ER: Estrogen receptor.

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  Discussion Top


Our previous laboratory work showed that during transplantation of ASC sheet on the damaged region of the rat uterus for over 30 days, epithelial cells could cover the excised uterine lumen and the ESC-like cells could be observed in the basal layer of the regenerated endometrium.[10] Sun et al. demonstrated that the soluble factors released by renal tubular epithelial cells (TECs) induced epithelial differentiation of hASCs, as evidenced by the significant elevation of the epithelial markers, downregulation of stem cell markers, and morphological changes in epithelial cells following culture in TEC-conditioned medium.[26] The aforementioned phenomenon suggests the essential role of the stem cell-based endometrium repair mechanism in thein vivo microenvironment, mainly referring to soluble factors secreted by the host cells. One of the beneficial effects might be the endometrial epithelial differentiation of ASCs. Therefore, to mimic thein vivo microenvironment, we cocultured hASCs and hEMCs in a transwell system, wherein the upper layer of the well was seeded with mature hEMCs and the bottom layer with hASCs. We then observed the changes in cell morphology and epithelial marker expression. In the transwell system, water and soluble molecules, except cells, could freely cross the membrane between the layers; hence, the outcome was due to the paracrine effect instead of immediate cell–cell contact. Our method of hEMC isolation was similar to that of Osteen et al. introduced in 1989.[32] According to their report, isolated populations of hEECs and hESCs exhibited over 95% homogeneity. Hence, substances produced by hEMCs play a key role in the endometrial differentiation of hASCs. The results of the present study showed that the shape of hASCs changed to a tadpole-like one following coculture with hEMCs. In addition to morphological changes, elevated expression of epithelial lineage markers, including CK18, CK7, CK19, and EMA, was noted, and this outcome became increasingly evident upon the addition of 17β-estradiol as well as TGF-β1, EGF, or PDGF-BB to the culture medium. The morphological changes as well as the cytokeratin and vimentin expression patterns of isolated hEMCs, were consistent with those reported previously.[32]

The human endometrium is a unique tissue that undergoes dramatic monthly remodeling during the menstrual cycle in preparation for an implanting conceptus. Remodeling is completed when the uterine cavity is totally covered by regularly structured EECs and ESCs. In this study, we selected adherent hEMCs as the coculture cell lines because we attempted to induce the differentiation of hASCs into hEECs. Considering that the stromal and epithelial cells engage in a synergistic manner during differentiation,[33] we did not further separate the two cell types via unit gravity sedimentation.[34] Suspension cells, such as neutrophils, macrophages, mast cells, and lymphocytes,[35] which are the predominant endometrial immune cells, were excluded from the transwell coculture system in the present study. Although the secretion function of neutrophils and macrophages might be partially responsible for angiogenesis in endometrial repair, proliferation, and differentiation,[36] their principal critical role is to disrupt the endometrial tissue during menstruation by secreting enzymes and cytokines such as matrix metalloproteinases, elastase, and interferon γ.[37] The lymphocytes chiefly function as immunomodulators to facilitate the success of embryo implantation and the subsequent development of the placenta and fetus rather than endometrial regeneration.[38] Whether lymphocytes can also induce epithelial differentiation of hASCs remains unclear and warrants further investigation.

However, with incomplete knowledge of factors involved in tissue regeneration, we cannot identify the key substance that initiates hASC differentiation. Several factors are released by hEMCs. TGF-β, EGF, insulin-like growth factor-1, basic fibroblast growth factor, and PDGF are the locally produced growth factors that participate in endometrial regeneration through autocrine and/or paracrine interactions between epithelial and stromal cells.[39],[40] Particularly, EGF, TGFβ, and PDGF-BB are required for candidate ESCs for both epithelial and stromal colony-forming activities,[35],[41],[42] indicating that these three factors are essential for stem cell differentiation. Felicia et al. verified that ERK–MAPK signaling is critical for adipogenesis, chondrogenesis, and adipogenesis of bone marrow-derived MSCs. Furthermore, PDGF signaling is critical for adipogenesis and chondrogenesis and TGF-β signaling for chondrogenesis. Accordingly, these three passages are sufficient for MSC growth in a serum-free medium.[43] Adipose- and bone marrow-derived MSCs share numerous characteristics. Thus, these three pathways might also work in the endometrial differentiation of hASCs. In the present study, the induction medium contained TGF-β1, PDGF-BB, and EGF. Following the induction culture without hEMCs, the expression of CK18 mRNA was elevated in Group 2 hASCs, although this change was not statistically significant. The result was partially because of the growth elements that were added in the medium.

Estrogen is important for endometrial remodeling. The coordinated and sequential actions of estrogen and progesterone direct major remodeling events, such as preparing a receptive endometrium for blastocyst implantation every month. Estrogen regulates endometrial cell survival, viability, and mitogenic effects via Esr1, which is the predominant endometrial estrogen receptor in mouse models.[44] Interestingly, the epithelial cells lack ER-α expression in the initial stage of endometrial repair in rhesus monkeys. ER-α could only be detected on the 3rd day of menstruation in rhesus monkeys.[45] Similarly, complete endometrial regeneration of all cellular components could occur without estrogen in a mouse model.[46] Thus, the neighboring ER-α–expressing niche cells are most likely to mediate the proliferation of epithelial cells. This hypothesis was proven by Chan and Gargett who demonstrated that Esr1 was expressed in a small population of mouse label-retaining cells, which is a group of candidate endometrial stem/progenitor cells with endometrial differentiation potential in mouse.[47] Unfortunately, whether human endometrial stem cells express ER-α remains unknown.[48] Moreover, there is no evidence to prove whether the role of endometrial repair is played by ESCs through a certain ER. Remarkably, in the present study, we found that both ER-α and ER-β were positively expressed in hASCs. Therefore, we added 17β-estradiol, which is the principal circulating estrogen in reproductive-age women, into the induction medium. Consequently, the mRNA expression levels of ER-α and ER-β in hASCs continually increased to the extent that hASCs differentiated into EECs, as attested by significantly elevated CK18 mRNA expression levels in the four groups. Thus, the endometrial differentiation of hASCs might be mediated by one of the ERs. Interestingly, the elevation of ER-β expression was statistically significant with the tense of the induction, indicating that ER-β is more likely a differentiation mediator.

Intracellular β-catenin is required by the Wnt canonical pathway, which is proved to be an important regulatory signaling axis that influences major developmental processes in the embryonic state and regulates maintenance, self-renewal, and differentiation of adult mammalian tissue stem cells.[49] Intracellular β-catenin can respond to Wnt ligand binding and act as a complex and dynamic signaling network and finally affect the fate of stem cells. In the present study, the expression of β-catenin mRNA elevated sequentially from Group 1 to Group 4, parallel with the tense of the induction, but the elevation did not reach the statistical significance. Our finding suggested that the β-catenin required canonical Wnt pathway might be a potential molecular mechanism participating in the process of endometrial epithelial differentiation of hASCs.

The association between estrogen and long noncoding RNA (lncRNA) H19 has been well studied in breast cancer progression. The proliferation, differentiation, and metastasis of ER-positive breast cancer cells induced by 17β-estradiol positively correlated with the H19 expression levels.[48] Cancer and stem cells are both self-renewing cells; however, for well-differentiated cells, this capacity eventually degrades. In our study, H19 expression significantly declined in hASCs following induction culture, when the differentiated cells became more mature and less self-renewable, indicating that H19 downregulation might be related to stem cell differentiation. Interestingly, among the four groups, the expression of H19 was significantly downregulated when the growth factors and 17β-estradiol were added. Confirming the most influencing factor for H19 expression remains difficult. Nonetheless, the growth factors we added (EGF, TGFβ, and PDGF-BB) were endogenous factors secreted by hEMCs.[39],[40] Meanwhile, 17β-estradiol was the exogenous substance required in endometrial remodeling. Therefore, estrogen is most likely associated with H19 expression, and this causality needs further evaluation.

In conclusion, hASCs could differentiate into EECs when cocultured with hEMCs, and this process was enhanced by the addition of EGF, TGFβ, PDGF-BB, and 17β-estradiol to the culture medium. This study, for the first time, showed ER-α and ER-β expression in hASCs and preliminary explored changes in ER-α, ER-β, β-catenin and H19 mRNA expression during hASC differentiation. However, the cross-talk between these pathways warrants further investigation. Moreover, the mRNA expression of H19 was negatively correlated with differentiation, which is seemingly related to the estrogen singling pathway, and the β-catenin required canonical Wnt pathway might be a potential molecular mechanism participating in the process of endometrial epithelial differentiation of hASCs. These evidences provide interesting viewpoints for future studies for exploring the mechanism of endometrial epithelial differentiation of hASCs.

Financial support and sponsorship

This work was funded by grants from the National Natural Science Foundation of China (No. 81671463) and the Key Research and Development Plan of Shaanxi Province (No. 2017ZDCXL-SF-02-03).

Conflicts of interest

There are no conflicts of interest.



 
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