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Biology of Reproduction, 2018, 99(5), 1057–1069
doi:10.1093/biolre/ioy121
Research Article
Advance Access Publication Date: 20 June 2018
Partial regeneration of uterine horns in rats
through adipose-derived stem cell sheets†
Huijun Sun‡ , Jie Lu‡ , Bo Li, Shuqiang Chen, Xifeng Xiao, Jun Wang,
Jingjing Wang and Xiaohong Wang∗
Department of Obstetrics and Gynecology, Tangdu Hospital, Fourth Military Medical University, 569 Xinsi Rd.,
Xian 710038, China
∗
Correspondence: Department of Obstetrics and Gynecology, Tangdu Hospital, Fourth Military Medical University,
Xinsi Road 569#, Xian 710038, China. Tel: +86-029-84717269; Fax: +86-029-84777690; E-mail: [email protected]
†
Grant Support: The project was supported by grants from the National Natural Science Foundation of China (81370710)
and the “Science and technology co-ordinating innovative engineering projects” of Science and Technology Department
of Shaanxi Province (2013KTCL03-07).
‡
These authors contributed equally to this work.
Edited by Dr. Romana Nowak, PhD, University of Illinois Urbana-Champaign.
Received 22 October 2017; Revised 24 February 2018; Accepted 19 June 2018
Abstract
Severe uterine damage and infection lead to intrauterine adhesions, which result in hypomenorrhea, amenorrhea and infertility. Cell sheet engineering has shown great promise in clinical
applications. Adipose-derived stem cells (ADSCs) are emerging as an alternative source of stem
cells for cell-based therapies. In the present study, we investigated the feasibility of applying
ADSCs as seed cells to form scaffold-free cell sheet. Data showed that ADSC sheets expressed
higher levels of FGF, Col I, TGFβ, and VEGF than ADSCs in suspension, while increased expression
of this gene set was associated with stemness, including Nanog, Oct4, and Sox2. We then investigated the therapeutic effects of 3D ADSCs sheet on regeneration in a rat model. We found that
ADSCs were mainly detected in the basal layer of the regenerating endometrium in the cell sheet
group at 21 days after transplantation. Additionally, some ADSCs differentiated into stromal-like
cells. Moreover, ADSC sheets transplanted into partially excised uteri promoted regeneration of
the endometrium cells, muscle cells and stimulated angiogenesis, and also resulted in better pregnancy outcomes. Therefore, ADSC sheet therapy shows considerable promise as a new treatment
for severe uterine damage.
Key words: uterine regeneration, cell sheet engineering, adipose-derived stem cells, cell differentiation.
Introduction
The human endometrium is a highly regenerative tissue that endures
cyclical episodes of proliferation, differentiation, and shedding more
than 400 times during a woman’s reproductive years [1, 2]. Two
layers compose the endometrium, the upper functionalis, which contains a large amount of glands, and the lower basalis, which is composed of branching glands and dense stroma [3]. The shedding of
endometrium functional layer is followed by sequential changes in
circulating sex steroid hormones during the menstrual cycle, and
the basalis remains during menstruation and from which the new
functionalis regenerates [4–6]. During the proliferative phase of the
menstrual cycle, the basal layer of the endometrium functions in cell
C Crown copyright 2018.
renewal and tissue regeneration, and the endometrium regenerates
during each cycle [7, 8]. Therefore, any endometrial dysfunction
occurs following damage to the basal layer [1, 9]. A variety of diseases or traumas are capable of causing severe uterine damage that
may lead to basal layer injury and, consequently, an inability of the
functional layer of the endometrium to regenerate, resulting in scar
formation [10, 11].
In severe cases, damage causes complete or partial scarring of
uterine cavity, also known as intrauterine adhesions (IUA), followed
by amenorrhea, infertility or abnormal placenta implantation, or
recurrent miscarriage [12–14]. Several therapeutic methods, such as
surgery, hormonal drugs, and cell therapy, have been adopted for
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Research Article
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Materials and methods
Rats
All experimental animals were provided by the Animal Center of The
Fourth Military Medical University and treated in accordance with
the guidelines of the Experimental Animals Management Committee
(Xian Province, China). Ethical approval was obtained from the
Ethics Committee of Military Medical Sciences (Shanxi, China) for
this study. Two-day-old female Sprague Dawley (SD) rats were used
for ADSC cell isolation. Six-to-eight-week-old female SD rats (250–
300g) were used to establish the model. SD rats were housed in the
temperature-controlled environment at 23 ± 3◦ C under the humidity
in the 44 ± 2% with a 12-h light and dark cycle. All the animals
access food and water ad libitum.
ADSC isolation and monolayer culture
Two-day-old female rats were selected and sacrificed by cervical dislocation to excise the adipose tissue from the inguinal subcutaneous
region (Supplementary Figure S1A). ADSCs were purified based on
a previously described protocol [33–35], isolated and cultured in
vitro. Briefly, after rinsing several times with phosphate-buffered
saline (PBS, Gibco, Carlsbad, CA), the tissue was minced into fine
pieces, digested with 0.75% collagenase for 60 min at 37◦ C, and
centrifuged at 800 r for 5 min. Adipocytes were discarded. Subsequently, adipose-derived stromal vascular fraction (SVF) was obtained, which was a mixed cell source contains endothelial cells,
immune cells, smooth muscle cells, stem cells, and other stromal
cells [36]. The SVF pellet was resuspended in PBS. The suspension
was filtered through an 80-μm nylon screen to remove tissue debris
and then centrifuged at 800 r for 10 min. The cell suspension was
incubated for 24 h in α-MEM (Gibco) supplemented with 10% fetal
bovine serum (FBS, Gibco) and penicillin/streptomycin (HyClone,
Logan, UT) at 38◦ C under humidified 5% CO2 conditions. After
72 h of culture, the cell medium was exchanged to remove nonadherent cells. The medium was then changed every other day, and
passaged cells were subcultured when they reached confluence (80–
100%). Third-generation cultured cells from a homogeneous cell
population were ultimately obtained and then labeled with CM-DiI
(CellTrackerTM CM-DiI; C7000, molecular probes, life technologies, Eugene, Oregon) (Supplementary Figure S1B). Cells at the same
passage in culture were used for all experiments.
Flow cytometric analysis
Rat ADSCs were recognized via immunophenotyping using monoclonal antibodies as previously described [37, 38]. Third-passage
ADSCs were identified by performing flow cytometry (FCM). Cells
were harvested via trypsin enzyme digestion (Gibco) and then centrifuged, washed and resuspended. Resuspended cells at a concentration of 106 cells/ml were incubated for 30 min on ice with
CD31 (abcam, Cambridge, UK), CD45 (abcam, Cambridge, UK),
CD29 (abcam, Cambridge, UK), CD44 (abcam, Cambridge, UK)
and CD90 (abcam, Cambridge, UK) monoclonal antibodies. The
distribution of mesenchymal stem cells (MSCs) was determined by
performing FCM (BD FACSVantage, USA).
Preparation of ADSCs sheet
Third-passage ADSCs were harvested for further experiments. To
construct cell sheets, 5 × 105 ADSCs were cultured in 6-well plates
for 14 days at 37◦ C in a humidified atmosphere with 5% CO2 .
The inducing medium consisted of α-MEM (Gibco), 10% FBS, 1%
antibiotic-antimycotic (HyClone), 10 nmol/L dexamethasone and
100 mg/ml ascorbic acid (Sigma). When the ADSCs sheet matured,
they were detached from the culture dish by scraping and floated
up in the medium as a membrane (Supplementary Figure S1C). The
ADSC sheets were reshaped from the circle to the rectangle and
layered according to uterine wounds in the rat model (Supplementary
Figure S1D).
Morphology of ADSCs sheet
The surface morphologies of the ADSC sheets and metal complexes were investigated by performing scanning electron micrograph (SEM) analysis [39]. For the SEM study, glass slides were
used as carriers for the cell sheets, which were rinsed twice with
PBS, fixed with 2.5% glutaraldehyde, dehydrated in step-wise manner in increasing concentrations of ethanol, and dried with a critical
point drier. The samples were then sputter-coated with platinum
and examined using a scanning electron microscope (Hitachi Model
S-3000N, Tokyo, Japan). For light microscopy studies, the samples were routinely processed, fixed in 4% formalin, dehydrated
and paraffin-embedded. Specimen morphology was observed based
on hematoxylin and eosin (H&E, Sigma) staining of conventional
paraffin sections.
Quantitative real-time PCR
Total RNA from the ADSCs sheet and ADSCs suspension was
isolated using TRlzol reagent (Invitrogen, MD, USA). Total RNA
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the treatment of endometrial fibrosis [12]. However, these strategies
have not shown much benefit [15, 16]. Effective treatments for severe
endometrial damage are limited. Therefore, tissue engineering may
represent a new method for the repair of uterine function.
Cell sheet engineering (CSE) is an innovative technology to regenerate injured or damaged tissues and has shown promising potential in the field of tissue regeneration. CSE involves cultured cells
on thermo-responsive culture dishes that form dense cell sheets that
detach when the temperature decreases. This technology was first developed by Okano in 1993 [17]. The primary benefit of this approach
is that it retains extracellular matrix (ECM) proteins, growth factors,
and larger amounts of cytokines without enzymolysis [18]. However,
the entire process has been criticized as relatively complicated, timeconsuming, and costly because thermo-responsive-culture dishes are
expensive [19, 20]. Thus, exploring novel approaches to improve
cell sheet construction is imperative. One feasible method involves
adding specific amounts of ascorbic acid to create cell sheets [19–22].
Ascorbic acid stimulates ECM production as well as DNA synthesis and subsequent cell proliferation [23–25]. Additionally, ascorbic
acid supplementation has diverse effects on stem cells, and it could be
used to maintain stem cell properties [26]. Ascorbic acid treatment
plays key roles both ∗∗ in enhances the generation of induced pluripotent stem cells and increasing adipose-derived stem cell (ADSC) proliferation and upregulating Oct4 and Sox2 expression [27].
ADSCs are a type of adult stem cell characterized by self-renewal,
multi-potential differentiation, immunosuppressive properties, and
low immunogenicity [28–30]. In addition to these advantages, ADSCs secrete trophic factors that enforce therapeutic and regenerative outcomes for a wide range of applications [31, 32]. ADSCs are
currently considered the most promising cell type in regenerative
medicine.
In this study, we sought to construct ADSCs sheet to evaluate the
therapeutic effects of cell sheets on regeneration in a rat model.
H. Sun et al., 2018, Vol. 99, No. 5
Regeneration of uterine horns in rats, 2018, Vol. 99, No. 5
Rat uterine horn damage model and ADSCs sheet
transplantation
To evaluate our animal model, vaginal smears were collected daily
between 08:00 and 10:00 AM for estrous cycle determination. Sixtysix uterine horns from 33 rats (66 uterine horns) were randomly
assigned to three groups after four estrous cycles: the ADSCs sheet
transplant group (cell sheet group) (n = 23 uterine horns), the control group (n = 23 uterine horns), and the sham group (n = 20
uterine horns). Adult SD rats were anesthetized (Supplementary
Figure S1E, first panel), and the uterine horns were exposed through
an abdominal incision. The uterus of rat was opened along the side
of nonmesometrium, and uterine walls comprise three layers—the
serosa, the myometrium and the endometrium, with the myometrium
consisting of circular smooth muscle near the lumen and subserosal
longitudinal smooth muscle (Supplementary Figure S1E, second
panel). A segment of circular smooth muscle and endometrium approximately 1.5 cm in length and 0.5 cm in width was excised from
both sides of the uterine horn, leaving the subserosal longitudinal
smooth muscle and the serosa intact (Supplementary Figure S1E,
third panel). Immediately after resection surgery, the resected surface
was pressed with hemostatic gauze for hemostasis, and the damaged
area was confirmed to be thin. In a self-correlative study, rat uterine side-by-side comparisons were observed for diversity restoration
effects between the cell sheet group and the control group. The cell
sheets show strong adhesion, of which we take advantage to complete the transplant operation. In the cell sheet transplant group
(the left uterine horn), triple-layer ADSCs sheet were implanted to
replace the segmental uterine excised tissue (Supplementary Figure
S1F), and the uterine surface wound with the ADSC sheets remained
exposed for 30 min to ensure engraftment. A 100-mm culture dish
covered the wound to prevent from water loss. The control group
(the right uterine horn) had uterine lesions that were left untreated
(Supplementary Figure S1F) and the sham group rats received abdominal incisions with the uterine horns intact. The uterine wound
was closed by 7-0 absorbable sutures and abdominal incision was
closed by 4-0 absorbable sutures. All rats received penicillin by intramuscular injection twice a day for three days. At 21, 30, and 60
days after surgery, all the uterine horns were tested by histological
and immunofluorescence stain (Supplementary Figure S1F).
Immunofluorescence analysis
Single-cell ADSC suspensions were labeled with CM-DiI
(CellTrackerT CM-DiI; C7000, molecular probes, life technologies,
Eugene, Oregon) according to the manufacturer’s instructions before
cell sheets were induced to trace the stem cells. Specimens from regenerating uteri were embedded in OCT (Leica, Nussloch, Germany),
frozen, and serially sectioned at a thickness of 6 μm with a Leica
cryostat (Leica, Nussloch, Germany). Frozen sections were incubated
with primary anti-estrogen receptor beta (ER, 1:200, ab3576, abcam, UK), anti-progesterone receptor (PR, 1:200, ab2756, abcam,
UK), anti-cytokeratin 18 (Ck, 1:200, ab133263, abcam, UK), anti-
Vimentin (Vm, 1:1000, ab92547, abcam, UK), alpha smooth muscle (Sm, 1:200, ab32575, abcam, UK), and CD31 (1:100, ab64543,
abcam, UK) antibodies at room temperature for 1 h in vivo. The sections were then washed three times with PBS, and FITC-conjugated
goat anti-rat IgG secondary antibodies (Alexa Fluor 488 Conjugate,
CST, USA) were applied for 30 min at room temperature. Additionally, all slides were incubated for 15 min with 4, 6-diamidino2-phenylindole (DAPI, Santa Cruz) for nuclear staining. Slides were
observed with a confocal laser scanning microscope (Leica, Nussloch, Germany). Three visual fields at 200× magnification were
randomly selected in each slide, and the average number of positive
vessels was determined to perform statistical analysis.
Histological analysis
Samples from in vivo tissues were fixed with 4% paraformaldehyde for 24 h at 4◦ C, then embedded in paraffin blocks and cut into
8-μm-thick tissue sections using previously described methods [41].
Changes in uterine tissue structure were observed by performing
H&E staining (Sigma), and the degree of fibrosis was evaluated with
a Trichrome Stain (Masson) Kit (Sigma) according to the manufacturer’s protocols. Staining results were analyzed using an optical
microscope (Leica, Nussloch, Germany).
Functional testing
Uterine function was assessed by determining whether the regenerating uterine horn was capable of implanting a fertilized ovum
and providing adequate nourishment to the developing fetus. Sixty
days postprocedure, male SD rats were cohabitated with females
(n = 13 uteri each in the cell sheet group and the control group, n =
10 uteri in the sham group). Vaginal plugs were tracked every day
at 8:00 AM to confirm whether mating had occurred. Animals were
euthanized 18.5 days after the appearance of a vaginal plug, and
uterine horns were examined to determine the number and position
of embryos.
Statistical analysis
Data were graphed and analyzed with GraphPad Prism 5 (USA). Statistical analysis of endometrial thickness, CD31 density, the mean
optical density of Sm-positive areas (AOD), and number of fetuses
per uterine were performed with one-way ANOVA. A number of
fetuses at scar site were calculated by independent sample t-tests between control and cell sheet group. All measurements are presented
as the mean ± SEM. Fisher’s exact test was performed for comparison of pregnancy rate at scar site in cell sheet group versus control
group. P-values < 0.05 were considered significant (∗ P < 0.05 or ∗∗ P
< 0.01).
Results
Multipotent rat ADSCs
The effects of enzyme digestion on phenotypic markers were assessed
(Figure 1A), and rat ADSCs strongly expressed the stem cell markers
CD29, CD44, and CD90 but were negative for CD31 and CD45
(Figure 1B–F). In addition, rat ADSCs demonstrated the potential
to differentiate into adipocytes and osteoblasts (Figure 1G and H).
CellTracker CM-DiI was used to mark and trace the growth of ADSCs that were transplanted into uteri (Figure 1I and J). The induction
efficiency (%) of CM-Dil into ADSCs was 100%.
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was applied for cDNA synthesis using an iTaq Universal One-Step
RT-qPCR Kit (Bio-Rad Laboratories, Hercules, USA). Quantitative
real-time PCR (qRT-PCR) was performed using Platinum SYBR
Green SuperMix (Invitrogen, USA) and an ABI Prism 7500 realtime PCR apparatus (Applied Biosystems). PCR primer pair (rat)
sequences are summarized in Supplementary Table S1. The PCR reactions were carried out as previously described [40]. GAPDH was
used as an internal control.
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ADSC sheet formation and transplantation
Scaffold-free ADSC sheets represent a new technology to regenerate
injured or damaged tissues. After 10–14 days of cultivation in inducing medium, ADSCs sheets were harvested from 6-well plates in
vitro. The first panel in Figure 2A shows the surface of a sheet that
detached at the bottom with the edge cocked, while the second panel
in Figure 2A was floating ADSCs sheet. The intact ADSC sheets were
harvested and reshaped to the rectangle. The reshaped ADSC sheets
were transplanted onto the damaged endometrium (Figure 2B). As
observed by cross-section H&E staining, the ADSCs were embedded in their own secreted ECM. Moreover, ADSCs sheets were composed of several layers of cells and abundant ECM as determined
by longitudinal section analysis (Figure 2C). Based on scanning electron microscopy examination, ADSCs sheets were compact overlayer
films. Distinct secretory granules displaying spherical vesicles were
present on ADSCs sheet surfaces (red arrow) (Figure 2D). Transforming growth factor β (TGFβ), collagen type I (Col I), vascular endothelial growth factor (VEGF), and fibroblast growth factor
(FGF) secretion from ADSC sheets significantly increased, meanwhile this ADSC sheets construct significantly increased the expressions of stemness genes such as Nanog, and Sox-2 but did not impact
gene Oct-4 expression compared to that in ADSCs in suspension
(Figure 2E).
Effects of the uterine resection model
The anatomical structures of the normal uterine area were obvious
and thick compared to those of the operative area which is observed by H&E and Masson’s trichrome staining. Based on H&E
staining, the uterus comprised the endometrium, muscle layers, and
serosa in normal areas. Abundant endometrial glands were observed
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Figure 1. Phenotype and multidifferentiation of rat ADSCs. (A) Rat ADSCs showed spindle-shaped fibroblastic morphology at the 3rd passage. (B–F) Flow
cytometric characterization of rat ADSCs. ADSCs were positive for CD29 (B), CD90 (C), and CD44 (D), but did not express CD45 (E) and CD31 (F). (G and H) ADSCs
were induced to differentiate into multiple lineages. (G) Rat ADSCs differentiated into adipocytes as indicated by cells stained with Oil red O. (H) ADSCs were
induced to differentiate into an osteogenic lineage as observed by Alizarin red S staining. (I and J) CM-DiI was applied to trace, locate and recognize rat ADSCs.
The appearance of labeled ADSCs under an optical microscope (I) and a fluorescent microscope (J).
Regeneration of uterine horns in rats, 2018, Vol. 99, No. 5
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in the endometrial stromal layer (Figure 3A, left panel). Masson’s
trichrome staining was performed on the uterine horns to characterize the extent of the muscle layers, which were composed of smooth
muscle cells in circular and longitudinal patterns in the nonoperative area, leaving behind only the longitudinal muscle layer in the
operative area (Figure 3A, right panel). Immunofluorescence staining with an anti-Ck antibody revealed Ck-positive epithelial cells on
the normal uterine adluminal surface and the endometrial gland in
the endometrial stromal layer (white arrow). No simple columnar
epithelial cells were found on the endometrial surface of the resected
tissue, and no endometrial glands were observed either (Figure 3B,
first panel). Vm-positive stromal cells were distributed in region
under the simple columnar epithelial cells (white arrow). Conversely,
we did not find Vm-positive stromal cells in the resected area of the
endometrium (Figure 3B, middle panel). Sm-positive muscle bundles
were labeled with green fluorescence following immunofluorescence
staining. As expected, no circular fibers were found in the muscle
layer. Only Sm-positive longitudinal muscle bundles were observed
in the resected area (yellow arrow). Conversely, Sm-positive circular
and longitudinal fibers were observed in normal areas (white arrow)
(Figure 3B, below panel). Based on these results, the endometrial
epithelial layer, basal layer, and circular fibers were successfully ablated in the uterine damage model, laying the foundation for the
regeneration of damaged uteri.
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Figure 2. Configuration of a 3D ADSCs sheet. (A) Visualization of the attachment and detachment of rat ADSCs sheet in vitro (bar: 5 mm). (B) H&E histological
staining of cross-sections and longitudinal sections of ADSCs sheet. (C) SEM images of ADSCs sheet formation and distinct secretory granules with the
appearance of spherical vesicles on cell sheet surfaces (red arrow). (D) Stemness and secretory cytokine gene expression by ADSCs sheet were detected by
qRT-PCR. Data are presented as the mean relative density ± SEM; ∗ P < 0.05, ∗∗ P < 0.01, compared to ADSCs (n = 3).
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Histological structures of regenerated uterine horns
Gross examination showed that luminal stenosis occurred in the surgical regions of the uteri in the control group at 30 days (Figure 4C),
and adhesions or pathological stenosis formed at 60 days (Figure 4E).
However, luminal stenosis and scars were not observed in the cell
sheet group (Figure 4D and F), and the uterine anatomy was similar
to the sham group (Figure 4A and B). Representative images show
that more blood vessels were present in the cell sheet group at both
30 and 60 days after surgery (Figure 4D and F), while the control
group exhibited pale serosa(Figure 4C and E).
To evaluate the effects of ADSC sheets on the regenerative recovery of uterine horns, H&E and Masson’s trichrome staining were
performed. As shown in Figure 4H, the cell sheet group exhibited
greater numbers of endometrial glands in the endometrial stromal
layer and more apparent neovascularization 30 days after surgery.
There were significant differences in endometrial thickness between
the cell sheet group (442.883 ± 28.185 μm) and the control group
(120.9433 ± 13.676 μm) (P = 0.000) (Figure 4J). At 60 days, the
specimens in the cell sheet group appeared near-normal and exhibited more angiogenesis. No epithelial-like cells were observed in uterine centers in the control group, but columnar epithelium lining was
present in the cell sheet group (Figure 4H). Endometrial thickness
in the cell sheet group (641.484 ± 58.284 μm) was significantly
greater than that in the control group (300.048 ± 44.903 μm)
(P = 0.000) but was thinner than the sham group (1044.226 ±
8.911 μm) (P = 0.000) (Figure 4K).
According to Masson’s trichrome staining on postoperative day
30, uterine tissue in the sham group was present in the muscle layer,
which consists of internal-circular muscle and external-longitudinal
muscle (Figure 4I). At 30 days, positively stained smooth muscle
cells turned light red and were arranged in a fascicular pattern in
the cell sheet group but not the control group (Figure 4I). At 60
days, we observed the orderly arrangement of smooth muscle cell
fascicles at near-normal levels in the cell sheet group. However, the
endometrium in the control group exhibited fibrous tissue hyperplasia and collagen deposition and eventually formed scars (Figure 4I).
Immunofluorescence staining of the regenerative
uterine horns
We assessed epithelial regeneration using an anti-Ck antibody. Ckpositive epithelial cells covered the surface of the excised uterine
lumen after 30 days in the cell sheet group. In contrast, few epithelial
cells were found at excision sites in the control group (Figure 5A).
The lumenal surface of the uterus was a simple columnar epithelium,
which became more organized and exhibited secretory glands after
60 days in the cell sheet group but lacked epithelial cells and secretory
glands in the control group (Figure 5A).
Uterine specimens from each group were tested for Vm expression by performing immunofluorescence microscopy to investigate
stromal cell regeneration. Vm-positive stromal cells in excised sections were detected by the presence of a bright green fluorescent
signal in the basal layer of the endometrium in the cell sheet group
(Figure 5B). Although Vm expression was also observed in the same
region in the control group, it was present at a lower frequency,
and the endometrium was thinner than that in the cell sheet group
(Figure 5B). At 60 days, greater numbers of endometrial glands were
observed in the endometrial stromal layer in the cell sheet group,
and the fluorescence intensities of Vm-positive stromal cells were enhanced compared with those at 30 days; however, the control group
still exhibited lower frequencies of Vm-positive cells (Figure 5B).
We used a panel of monoclonal rabbit anti-rat antibodies raised
against smooth muscle actin (Sm) to explore smooth muscle regeneration at sites of injury. At 30 days, new circular muscle bundles formed a thin successive layer beneath the endometrium in the
cell sheet group. However, muscle bundles were absent at injured
sites in the control group with the exception of anastomosis sites
(Figure 5C). AOD in the cell sheet group (0.035 ± 0.003) was higher
than that in the control group (0.010 ± 0.001) (P = 0.000) but
lower than that in the sham group (0.074 ± 0.003) (P = 0.000)
(Figure 5E). At 60 days, some collagen deposition and scar tissue
formation was observed in the control group, but no obvious circular muscle bundles had formed. The smooth muscle tissue converged
to form thick muscle layers in the cell sheet group that were denser
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Figure 3. Experimental procedure for the rat uterine regeneration model. (A) Schematic illustration showed the experimental procedure. (B) Uterine horns were
exposed, and half of the uterus, approximately 1.5 cm in length and 0.5 cm in width, was resected and removed. Triple-layer ADSCs sheets were transplanted
onto the damaged inner uterine surface, and the uterine wound was left open for 30 min after transplantation to ensure engraftment. (C and D) To assess
the uterine damage model, uteri were resected, and specimens were subjected to histological analysis after surgery. (C) Specimens stained with H&E and
Masson’s trichrome stain were observed microscopically. Black arrows and blue arrows indicate endometrial glands and the circular muscle layer, respectively.
(D) Immunofluorescence staining of specimen components using antibodies against Ck, Vm, and Sm. White arrows show the endometrial epithelial layer (first
panel), endometrial basal layer (middle panel), and circular muscle layer, respectively (last panel). Yellow arrows indicate the presence only of the remaining
longitudinal muscle layer in the regeneration model. Dotted lines indicate repair sites.
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Figure 4. Gross view and histological evidence for uterine horn regeneration among the groups. (A–F) Gross observations of surgical regions in the lower
segment of the uterus at 30 days (A–D) and 60 days (E, F) in the sham group (A, B), control group (C, E), and cell sheet group (D, F). The appearance of surgical
regions indicated superior vascularization for the repair of segmental defects in the rat uterus in the cell sheet group compared with the control group. H&E
(H) and Masson’s trichrome staining (I) of the uterine horns after 30 or 60 days post-treatment in the sham group, control group, and cell sheet group. (J, K)
Statistical analysis of endometrial thickness revealed a thicker endometrium in the cell sheet group than in the control group; however, the endometrium was
thinner than that of the sham group. Dotted lines indicate repair sites. Black arrows indicate endometrial glands in the basal layer. Data are presented as the
mean ± SEM. ∗ P < 0.05 and ∗∗ P < 0.01.
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Figure 5. Immunofluorescence staining indicated Ck (A), Vm (B), Sm (C), and CD31(D) levels in each group at 30 or 60 days after surgery. (E) We assessed AOD
using IPP software and observed the continuous improvement of muscle bundle status over time. (D) CD31 expression, indicative of neovascularization in the
regenerated endometrium, was detected at 30 and 60 days in the sham group, control group, and cell sheet group. (F) Based on a statistical analysis of capillary
vessel number from at least three randomly selected fields at a magnification of 200×, cell sheet promoted blood vessel regeneration. Each experiment was
repeated four times. Dotted lines indicate repair sites. Data are presented as the mean ± SEM. ∗ P < 0.05 and ∗∗ P < 0.01.
Regeneration of uterine horns in rats, 2018, Vol. 99, No. 5
ADSC differentiation in vivo
Third-passage ADSCs were incubated with CellTracker CM-DiI and
exhibited red fluorescence (Figure 1J). Twenty-one days later, ADSCs fused with host cells and prompted local cells migration to the
wound edges. Notably, clusters of cells labeled with red fluorescence were primarily distributed in the basal layer of the regenerated endometrium (Figure 6A). Based on further observations of
uterine sections, some ADSCs may have directly differentiated into
Vm-positive endometrial stromal cells (Figure 6D), while others expressed ER (Figure 6B) and PR (Figure 6C), which are also expressed
in endometrial stromal cells. These typical images illustrate the differentiation of ADSC sheets during development in the uterine microenvironment.
Functionality of regenerated uterine horns
To investigate the effects of cell sheet treatment on the functional
recovery of regenerated uterine horns, the ability to accept an embryo was tested. At 60 days, female rats were mated with male rats
(the female: male ratio was 2:1) and were checked for pregnancy at
18.5 days after the appearance of the pessary. Fetuses implanted in
all of the uterine horns in the sham group (Figure 7A and B). In the
control group, embryos were rarely observed at scar sites, but the
majority implanted at normal sites (Figure 7C). In contrast, fetuses
were observed both in regenerated uterine tissue and in normal uterine tissue in the cell sheet group (Figure 7D). Pregnancy rate at the
scar site was 69.23% for the cell sheet group and 15.38% for the
control group (P = 0.005) (Table 1). In addition, the total number
of implanted embryos in the control group (2.308 ± 0.634) was significantly reduced compared with that in the cell sheet group (5.230
± 0.975) (P = 0.020) (Table 1). Furthermore, the number of embryos that implanted at the scar site was noted. In terms of scar site
pregnancy, 0.846 ± 0.222 embryos demonstrated successful nidation, but in the control group only 0.154 ± 0.104 embryos showed
successful scar site implantation (P = 0.009) (Table 1). Thus, cell
sheets grafted onto injured endometrium in vivo reestablished maternal uterine functionality to support embryo implantation and fetal
development.
Discussion
The clinical application of cell sheet therapy has been reported for
the cornea [42], liver [43, 44], arteries [45], esophagus [46], bone
[47], heart [48, 49], and brain [50]. Here, we revealed the feasibility
of partial resection of the rat uterus to create a uterine regeneration
model. This method permitted a transplantable ADSC sheet to firmly
attach to a site of injury to maintain lumenal architecture. We also
demonstrated remodeling of the 3D architecture of the rat uterus
via engraftment of an ADSC sheet onto a partially excised uterus.
Muscle bundles were rebuilt, and columnar epithelial-like cells were
reconstituted. Furthermore, ADSCs were capable of differentiating
into stromal-like cells when ADSC sheets were transplanted into the
uterus.
In regenerative medicine, other approaches have been reported
to effectively regenerate injured uterine tissue, such as recellularization of a decellularized uterine matrix [51], collagen scaffolds
in combination with collagen-binding human basic growth factor
[52, 53] or the application of bone marrow MSCs [21]. However,
inflammatory responses are induced by scaffold materials during
biodegradation in vivo, and the resultant constructs exhibit low cell
density [54]. The 3D tissue-like structures of cell sheets are formed
by cell-to-cell junctions and deposited ECM in the absence of any
scaffolds. Therefore, an immune response is not initiated, and inflammatory responses do not accompany transplantation [55]. Cell
sheets are also easily shaped into different structures for different applications and are especially suitable for the narrow uterine
lumen.
With their capacity for self-renewal, multidirectional differentiation and low immunogenicity, ADSCs are emerging as an alternative
stem cell source for cell-based therapies [56]. A previous study described the treatment of severe full-thickness uterine injury in rats
using bone marrow MSCs for collagen scaffold transplantation [36].
MSCs primarily localized to the basal layer, and a small number differentiated into stromal cells. In the present study, we also observed
the random distribution of most ADSCs onto the basal layer. Meanwhile, ADSCs were capable of differentiating into stromal cells. This
preliminary analysis illustrates a possible mechanism to explain the
effects of ADSC sheets on the injured uterus.
The basal layer of the endometrium contains mesenchymal stemlike cells that express markers similar to bone marrow-derived stem
cells, characterized by multiple differentiation potential [57]. Therefore, a new functional layer regenerates during each menstrual cycle
[58]. Uterine endometrial cells adjacent to the wound margin have
the potential to regenerate the wound site via uterine cell proliferation and migration. However, according to our study, uterine
regeneration was insufficient in the control group. Therefore, the
3D structure of the ADSC sheets provided an efficient cell microenvironment that was critical for ADSC differentiation and uterine
cell regeneration. In the present study, ADSC sheets demonstrated
higher FGF, Col I, TGFβ, and VEGF expression compared to that
in ADSCs in suspension. These bioactive factors not only promote
growth and development but also inhibit apoptosis, benefit local
cell proliferation, and are conducive to ADSC differentiation [36,
59, 60]. TGFβ is a pleiotropic cytokine that is important in the
regulation of joint homeostasis and disease. TGFβ also regulates
many of the processes common to cell differentiation, tissue repair and inflammation [61]. Angiogenesis, which is primarily mediated by the VEGF pathway, is crucial for tube formation and cell
proliferation [62]. According to previous studies, collagen-binding
VEGF remodels full-thickness uterine injury [53]. bFGF exerts many
Downloaded from https://academic.oup.com/biolreprod/article-abstract/99/5/1057/5040762 by Indian Institute of Technology Kharagpur user on 17 August 2019
and arranged. However, at the site of anastomosis, only circular
fibers were observed, rather than circular and longitudinal fibers
(Figure 5C). The AOD in the cell sheet group (0.059 ± 0.031) was
still significantly higher than the control group (0.022 ± 0.036)
(P = 0.000), but no statistically significant difference was observed
between the cell sheet group and the sham group (0.065 ± 0.035)
(P = 0.341) (Figure 5E). Thus, cell sheets promoted the regeneration
of injured muscle bundles, and this outcome became increasingly evident over time. Neovascularization is one of significant standards to
evaluate the regeneration of the injured uterine regions. At 30 days
after the implantation of cell sheet grafts, blood vessel density within
the cell sheet group was higher, with neovascularization located near
the site of injury (Figure 5D). Blood vessel density in injured uterine
regions in cell sheets group (26.40 ± 1.806) was significantly higher
than that in the control group (15.80 ± 2.634) (P = 0.011), and
less than the sham group (54.40 ± 4.130) (P ≤ 0.001) (Figure 5F).
At 60 days, the blood vessel was increased in all groups compared
with 30 days, however, the cell sheet group (42.80 ± 3.262) showed
a significant increase compared to control group (24.40 ± 2.112)
(P = 0.002) and still less than the sham group (54.40 ± 2.227) (P =
0.019) (Figure 5F).
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H. Sun et al., 2018, Vol. 99, No. 5
biological functions, such as stimulating the proliferation of fibroblasts, skeletal myoblasts, vascular endothelial and smooth muscle
cells, and is also essential for cell survival, migration, and matrix
production/degradation [63]. As reported by Li, collagen scaffolds
loaded with collagen-binding FGF effectively repaired damaged endometrium through angiogenesis and epithelial regeneration [52].
Wound-healing synthesis processes, which include the rapid deposition of ECM, may be regulated by the ECM microenvironment
[64]. A variety of components constitute the ECM, but the defining component is Col I [65]. After the transplantation of ADSC
sheets into the injured uterus, different growth factors and extracellular vesicles interact to prompt wound healing and mediate cell
regeneration.
Our objective in evaluating regenerated uterine function was to
assess pregnancy potential. In the present study, pregnancy rates at
the scar site were 69.23% in the cell sheet group and 15.38% in the
control group. In addition, the total number of implanted embryos
in the control group was significantly reduced compared with that
in the cell sheet group. Embryos were rarely observed in scar sites
in the control group, but fetuses were observed in both the regenerated uterine tissue and the normal uterine tissue in the cell sheet
group. Therefore, this therapeutic method not only remodeled the
structure of the injured uterus by recovering the lumen-adjacent epithelial layer, basal layer, and smooth muscle layer but also restored
functionality as evidenced by successful pregnancy.
Conclusions
The present study reviewed a novel approach for the regeneration of
partial uterine defects. Thus far, tests have only been conducted on
animals in a laboratory setting, but human testing is anticipated for
the next phase. However, many challenges remain. Cell sheet therapy
may be a new approach for treating partial uterine defects due to
dilatation and curettage after abortion or postpartum, myomectomy
for uterine fibroids, adenomyomectomy for uterine adenomyosis,
severe IUA from endometrial tuberculosis, and conization in uterine
cervical cancer. Our group is conducting further experiments with
human subjects to benefit mankind.
Downloaded from https://academic.oup.com/biolreprod/article-abstract/99/5/1057/5040762 by Indian Institute of Technology Kharagpur user on 17 August 2019
Figure 6. The location and differentiation of ADSCs in the newly regenerated uterine endometrium 21 days after cell sheet transplantation as determined by
immunofluorescence staining. (A) ADSCs, labeled red, were tracked using a confocal microscope and were primarily located near the basal membrane of the
regenerated rat endometrium. Representative fluorescent images demonstrate positive, ER (green) (B), PR (green) (C), and Vm (green) (D) protein expression
(arrows) in some ADSCs after the transplantation of labeled cell sheets, indicating ADSCs may have differentiated into endometrial epithelial cells and stromal
cells. ADSCs labeled with CM-DiI (red) were tracked using a confocal microscope, and cell nuclei were stained with DAPI (blue). Dotted lines indicate repair sites.
Presence of double positive cell is indicated by white arrow.
Regeneration of uterine horns in rats, 2018, Vol. 99, No. 5
1067
Table 1. Comparison of reproductive outcomes among different groups with regenerated uterine horns. Data are mean ± SEM or n (%).
Pregnancy rate at scar site
Number of fetuses per uterine horn: mean ± SEM
Number of fetuses at scar site: mean ± SEM
Sham (n = 10)
Control (n = 13)
Cell sheet (n = 13)
8.100 ± 0.314
2 (15.38%)
2.308 ± 0.634b
0.154 ± 0.104
9 (69.23%)a
5.230 ± 0.975b,c
0.846 ± 0.222d
P < 0.01, vs. control group.
P < 0.05, vs. sham group.
c
P < 0.05, vs. control group.
d
P < 0.01, vs. control group.
a
b
Supplementary data
Supplementary data are available at BIOLRE online.
Supplementary Figure S1. Schematic illustration of experimental procedure. (A) Adipose tissues were resected from 2-day-old female rats.
(B) The ADSCs were isolated from the tissue with enzymolysis, and
then labeled with CM-DiI. (C) ADSCs were cultured with induced
medium for 14 days at 37◦ C in a humidified atmosphere with 5%
CO2 . (D) ADSCs sheet was harvested from the insert, reshaped, and
layered according to the target tissue site. (E) A segment of tissue
approximately 1.5 cm in length and 0.5 cm in width was excised
from both sides of the SD rat uterine horn. (F) In the cell group
(the left uterine horn), triple-layer cell sheets were transplanted on
the damaged inner uterine surface. In the control group (the right
uterine horn), the uterine wound left untreated. At 21, 30, and 60
days after surgery, all the uterine horns were tested by histological
and immunofluorescence stain.
Supplementary Table S1. qPCR primers of target genes.
Acknowledgments
The authors thank Meng Xu for assistance in immunofluorescence and Chong
Lu forhelp in flow cytometry. We also acknowledge the technical assistance
from the staff of the Key Laboratory of Brain Stress and Behavior of Tangdu
Hospital, The Fourth Military Medical University.
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