Female Sprague-Dawley (SD) rats (8-weeks-old; 200–250 g) were used for this study, and were housed in a climate-controlled facility with free access to food and water. All experiments were approved by the Institutional Animal Care and Use Committee of the Shanghai Jiao Tong University, and were consistent with the NIH Guide for the Care and Use of Laboratory Animals (NIH Publications No. 80-23).

Bone marrow samples were isolated by flushing the femurs, tibiae, and humeri of C57BL/6 mice (female, 8 weeks old) with cold PBS. The bone marrow cells were passed through 70 um mesh filters (BD Biosciences, United States) and were then cultured in DMEM containing 10% fetal bovine serum (FBS, Gibco, MA, United States) and 1% streptomycin/penicillin (Sigma, MO, United States). BMSCs from passage 3 were used for downstream experiments.

A murine Mesenchymal Stem Cell Functional Identification Kit (Cyagen, SuZhou, China) was used to evaluate the ability of isolated BMSCs to undergo in vitro differentiation. Briefly, BMSCs were added to 6-well plates (10⁵ cells/well) in media containing adipogenic or osteogenic differentiation medium once 70–80% confluent. Supernatants were replaced every 2–3 days. On day 21, Alizarin Red staining and Oil Red O staining were used to assess osteoblastic and adipogenic differentiation, respectively (Liu et al., 2020). Cells were imaged via fluorescence microscopy (Olympus, Tokyo, Japan).

An adenoviral vector capable of promoting stable CTF1 overexpression was initially constructed. Briefly, the CTF1 full-length gene was cloned via RT-PCR, with the GFP-encoding pDC316-CMV construct being utilized as a shuttle vector. Following nuclease digestion, the sequence of the pDC316-CTF1 construct was confirmed. The adenovirus packaging plasmid (pBHGlox_E1.3Cre) was then co-transfected into HEK293 cells along with this construct, yielding the GFP-CTF1 adenoviral vector encoding CTF1, as confirmed via PCR. Following purification, amplification, and titration, this GFP-CTF1 adenovirus was used to infect BMSCs, after which CTF1 expression levels were confirmed via qPCR and Western blotting. An adenoviral vector encoding GFP served as a negative control construct. Infections were conducted for 24 or 48 h at multiplicity of infection (MOI) values of 250, 500, and 1,000 in order to optimize transduction efficiency.

Exosomes isolated from BMSCs or CTF1-BMSCs were respectively referred to as BMSCs-exo and C-BMSCs-exo. Briefly, once cells were 90% confluent, the overlying culture medium was discarded, and cells were rinsed once with PBS, after which media containing 10% exosome-depleted FBS and penicillin/streptomycin was added and cells were cultured for 48 h. Conditioned media was then collected and used to isolate the exosomes therein. Briefly, media was first centrifuged for 10 min at 400 × g to remove remaining cells, after which debris was removed by spinning supernatants for 20 min at 2,000 × g at 4°C. Samples were then centrifuged for 40 min at 15,000 × g, and supernatants were passed through a 0.22-μm filter (Millipore, MA, United States) prior to ultracentrifugation for 70 min at 120,000 × g. Exosomes were then washed with PBS and this ultracentrifugation step was repeated to isolate exosomes. Samples were maintained at or above 4°C at all times. A BCA Protein Assay Kit (Beyotime, Shanghai, China) was used based on provided directions to assess exosome protein concentrations, after which these particles were characterized via Western blotting, Nanoparticle Tracking Analysis (NTA), and transmission electron microscopy (TEM) (Wang et al., 2019).

All rats were anesthetized by intraperitoneal injection of 2% sodium pentobarbital (0.3 ml/100 g), and the uterine horns were exposed via an abdominal incision. The proximal and distal portions of the uterus were then gently clamped with two vascular clamps, after which 95% ethanol was injected into the uterine cavity for 60 s. Vascular clamps were then released, gently squeezed to remove residual ethanol, and damaged sites were washed with PBS thrice to further remove remaining ethanol, as discussed in prior reports (Lin et al., 2021). HyStem-HP hydrogel (Catalog: GS315, Glycosan Biosystems, UT, United States) was utilized as a carrier for exosome delivery. This hydrogel was selected as it is composed of thiol-modified hyaluronan, HA, and thiol-modified heparin, and can undergo cross-linking in situ upon injection (Qin et al., 2016; Zhang et al., 2019). A total of 80 rats (n = 6–8 rats per group from three independent experiments) were assigned at random to four treatment groups (n = 20 for each group, 10 were sacrificed for HE, Masson and immunofluorescent evaluation and the other 10 for pregnancy test): 1) a sham-operated control group in which the uterine horn was injected with PBS (200 μl); 2) a model group in which the uterine horn was injected with 95% ethanol as in prior studies (Lin et al., 2021); 3) a model group that was treated with BMSCs-exo (100 μg/ml) in a hydrogel suspension via injection into the uterine horn (20 μg in 200 μl hydrogel); and 4) a model group that was treated with C-BMSCs-exo (100 μg/ml) in a hydrogel suspension that was injected into the uterine horn (20 μg in 200 μl hydrogel). For both exosome-treated groups, exosomal suspensions were administered 30 min after ethanol-mediated damage was induced. When three estrus cycles had passed after hydrogel injection, rats were euthanized and samples of uterine tissue were collected and frozen or sectioned for downstream analyses.

For wound healing assays, human umbilical vein endothelial cells (HUVECs; ATCC) (4 × 10⁴/well) were added to the lower chamber of a well containing a Transwell assay insert (8-μm pores, Corning, United States), with 20 μg of BMSCs-exo or C-BMSCs-exo being added into the upper compartment. The monolayer was scratched with a sterile pipette tip, washed with PBS, and cells were then cultured for 24 h in high-glucose DMEM, with cells being imaged at 0, 12, and 24 h via inverted microscope (Olympus Microscopes, Tokyo, Japan). Wound closure in six randomly selected fields of view was then quantified using Photoshop (Adobe Systems Inc., CA, United States), with data being normalized to mean values in the control group.

For migration assays, HUVECs (4 × 10⁴) were instead added to the upper chamber of the Transwell insert, with BMSCs-exo or C-BMSCs-exo (20 μg) being added in the lower chamber. Following incubation for 18 h at 37°C, cells that had migrated to the lower chamber were fixed, stained with 0.5% crystal violet, and quantified via microscopy.

HUVEC proliferation was assessed via immunofluorescent Ki67 staining. Briefly, cells were fixed using 4% paraformaldehyde, permeabilized using 0.1% Triton X-100, and stained using anti-Ki67 (1:200; Abcam), followed by staining with an appropriate secondary antibody conjugated to AF488 (Abcam). Nuclear counterstaining was then performed, and Ki67-positive cells were quantified using the ImageJ program.

To assesses the impact of BMSCs-exo or C-BMSCs-exo on endothelial tube formation, HUVECs (1 × 10⁴/well) were plated in 24-well plates on growth factor-reduced Matrigel (BD Biosciences, United States) for 16 h in the presence of appropriate exosomes, after which tubes were imaged via microscopy and randomly chosen pictures were analyzed with the Image J Angiogenesis Analyzer function.

Embryo implantation was measured as a correlate for endometrial receptivity. Briefly, rat estrus cycle progression was assessed via a vaginal smear approach. At 8 weeks post-surgery, female rats (10 rats, 20 uterine horns per treatment group) were mated with sexually mature male rats at a 1:1 ratio on the day of estrus, with rats being examined the following morning for the presence of a vaginal plug. On day 18 after the observation of a vaginal plug, rats were euthanized, and the size, number, and weight of fetuses therein were assessed, as were the locations of these fetuses.

Endometrial tissue sections were fixed overnight in 4% paraformaldehyde, after which they were embedded in paraffin blocks and cut into 8-μm-thick tissue sections using previously described methods (Hashimoto et al., 2016). Sections were then stained with hematoxylin and eosin (H&E) or were subjected to Masson’s trichrome staining based upon provided protocols. Endometrial sections were additionally stained for CD31 and α-SMA as detailed previously (Qin et al., 2013), with DAPI as a nuclear counterstain.

Total protein was isolated from the HUVECs co-cultured with 100 μg/ml BMSC-exos and 100 μg/ml CTF1-BMSC-exos for 48 h in vitro, and endometrial tissues of the BMSC-exo treatment group and the C-BMSC-exo treatment group in vivo, after which protein levels in the prepared lysates were measured. Protein was then separated via SDS-PAGE, transferred to PVDF membranes, and the resultant blots were probed overnight with antibodies specific for mTOR, p-mTOR, JAK, p-JK, PI3K, p-PI3K, STAT3, p-STAT3, AKT, p-AKT, or β-actin (1:500, Abcam) at 4°C. Relative protein band density was then assessed with an Odyssey infrared imaging system (LI-COR, NE, United States). The details of the quantification process of density values are specified in Image J software. Quantification of protein was normalized to β-actin.

Isolated BMSCs were characterized by staining them with antibodies specific for CD29 (Biolegend), CD34 (eBioscience), CD44 (Biolegend), CD45 (Biolegend), and CD90 (Biolegend) in a 100 μl volume of FACS buffer (Sigma, CA, United States). After two washes with FACS buffer, cells were fixed in Cytofix (BD Pharminigen, CA, United States) and analyzed with an ACEA NovoCyte flow cytometer. The FlowJo software was used for all data analyses.

Data are means ± SD and were analyzed via one-way ANOVAs with Tukey’s multiple comparison test using SPSS 22.0. p < 0.05 served as the significance threshold. In all figure legends, “n” represents the number of samples or rats utilized in the indicated experiments and technical and biological replicates are specified for each experiment.

First, we isolated BMSCs from C57BL/6 mice and culturing these cells for three passages, after which cells were used for downstream experiments (Figure 1A). Flow cytometry confirmed that these cells were negative for hematopoietic markers (CD34) and were positive for BMSC markers antigens (CD29, CD90, CD44, and CD45) (Figure 1B). Consistent with their identity as multipotent progenitor cells, these BMSCs were able to differentiate into osteoblast- and adipocyte-like cells when treated with appropriate differentiation media, as confirmed via Alizarin Red staining and Oil Red O staining, respectively (Figure 1C). BMSCs-derived exosome were then collected via differential centrifugation from BMSC-condition media. TEM analyses confirmed these BMSCs-exo to exhibit rounded or cup-shaped morphologic characteristics consistent with those of exosomes (Figure 1D), while TRPS measurements indicated these exosomes to be 60–120 nm in size (Figure 1E). Western blotting also revealed these particles to be positive for exosomal surface markers including Alix, CD9, CD63, and CD81 (Figure 1F). Together, these data confirmed that we had successfully enriched mouse BMSCs-derived exosome.

An adenoviral vector was used to stably overexpress CTF1-GFP in murine BMSCs as detailed in the Materials and Methods section, with GFP overexpression serving as a negative control. A range of adenoviral transduction intervals and MOI values were tested to maximize the efficiency of this overexpression approach (Figure 2).

An adenoviral vector was used to transduce CTF1 into BMCSc at an MOI of 500, with maximal transduction efficiency being evident after 48 h (Figure 2). The transduction ratio did not vary markedly between CTF1 and GFP in BMSC cells (Figure 2A). Western blotting and PCR confirmed that these CTF1-transduced BCSCs exhibited significant increases in CTF1 expression, whereas it was almost undetectable in the control BMSC group (p < −0.05) (Figures 2B–D).

Angiogenesis is an essential process whereby stem cell-derived exosomes can drive endometrial regeneration. As such, we next evaluated the effects of C-BMSC-exo treatment on the proliferation, migration, and tube formation activity of HUVECs in vitro. In a wound healing assay, C-BMSC-exo treatment was associated with significant improvements in wound closure at 12 and 24 h time points relative to BMSC-exo treatment (p < 0.05; Figures 3A,B). In migration assays, C-BMSC-exo treatment was associated with significant increases in cellular migration relative to BMSC-exo treatment (99 ± 4 vs. 247 ± 4 cells/field; p < 0.05) (Figures 3A,C). These data thus suggested that CTF1 overexpression was linked to enhanced migratory activity upon BMSC-exo treatment.

In proliferation assays, Ki67 staining was performed following a 3-day treatment, revealing that the majority of cells in both the BMSC-exo and C-BMSC-exo groups were Ki67-positive (Figure 4A), with C-BMSC-exo treatment being associated with greater positivity relative to BMSC-exo treatment (Figure 4C). These results thus indicated that C-BMSC-exo treatment was sufficient to promote sustained in vitro endothelial cell proliferation.

Tube formation assays were next conducted to evaluate the angiogenic potential of treatment with these different exosomal preparations. In line with the above data, we found that C-BMSC-exo treatment was associated with significantly enhanced tube formation relative to BMSC-exo treatment (Figures 4B,D). Overall, these findings suggested that CTF1 overexpression led to enhanced proangiogenic activity following BMSC-exo treatment.

We next employed an intrauterine anhydrous ethanol injection approach to model endometrial damage in vivo in rats. Endometrial damage was assessed in these animals via a series of histological approaches after three estrus cycles had passed following damage. Endometrial damage and thinning were evident in model group rats such that in some cases the endometrium and myometrium could not be distinguished from one another. Significant endometrial regeneration was evidence in rats which had been injected with C-BMSC-exo preparations, as evidenced by increased angiogenesis and areas of well-distributed cells (Figure 5A). Such treatment was also associated with appropriately structured stromal tissues, epithelial layers, and secretory glands. Rats in the sham-operated control, model, BMSC-exo group and C-BMSC-exo group exhibited endometrial tissues with a thickness of 490, 200, 350, and 425 µm, respectively, underscoring the potent regenerative effects of C-BMSCs-exo treatment (Figure 5B).

Masson’s trichrome staining was next conducted to assess collagen deposition within uterine tissue sections from these treated rats, revealing significant reductions in endometrial fibrosis and enhanced muscle fiber growth following BMSCs-exo treatment (Figure 5C). Importantly, C-BMSCs-exo treatment was associated with even more robust anti-fibrotic activity (Figure 5C), suggesting that C-BMSCs-exo can effectively prevent tissue damage and fibrosis, thus promoting efficient vascular and glandular proliferation following the induction of endometrial damage.

Endometrial regeneration is dependent on angiogenesis, which facilitates oxygen and nutrient delivery to damaged tissues (Gurtner et al., 2008). As such, we next explored the effects of C-BMSCs-exo treatment on in vivo neovascularization (Figure 6). We therefore stained endometrial tissues with the anti-CD31 to detect endothelial cells, revealing that there were significantly higher levels of CD31 staining in the BMSCs-exo and C-BMSCs-exo treatment groups as compared to the model group (Figure 6A), C-BMSCs-exo treatment being associated with the highest levels of staining. Immunofluorescent staining results further confirmed that C-BMSCs-exo treatment resulted in significantly increased in vivo neovascularization (Figure 6C).

Anti-α-SMA was next used to stain tissues in order to assess myometrial regeneration (Figure 6B). Animals in the control and BMSCs-exo groups exhibited a thin, continuous layer of circular muscle fibers, whereas this was not the case in the model groups. Rats in the C-BMSCs-exo group exhibited a significant increase in the number of muscle bundles as compared to the BMSCs-exo group (Figure 6B). The α-SMA staining area in the C-BMSC-exo group was significantly higher than that in the BMSCs-exo group (Figure 6D). Together, these data indicated that BMSCs-exo treatment promoted myometrial regeneration.

To further evaluate the functional effects of C-BMSC-exo treatment on endometrial regeneration, we next measured implantation rates in different treatment groups by counting embryos in rats on gestational day 18 (Figure 7; Supplementary Table S1). Sham-operated control rats exhibited the highest implantation rates, but the rates in the C-BMSC-exo treatment group were significantly higher than those in the untreated and BMSC-exo groups (Figure 7), suggesting the ability of C-BMSC-exo treatment to restore fertility by promoting functional endometrial regeneration.

Lastly, we performed a series of Western blotting analyses examining the signaling pathways whereby CTF1 may mediate beneficial fertility outcomes. We found that C-BMSCs-exo groups exhibited significantly higher levels of JAK/PI3K/mTOR/STAT3 phosphorylation relative to BMSCs-exo control groups (Figures 8A,C), with these increases being confirmed via statistical analyses (Figures 8B,D). And no change in AKT expression was observed in the C-BMSCs-exo groups relative to the corresponding BMSC-exo controls. We speculated that STAT3 was a key signaling mediator in this context. Indeed, p-STAT3 has previously been reported as a regulator of VEGFR2 activity (Chen et al., 2008). CTF1 can promote angiogenesis and myocardial cell survival through the induction of VEGF-dependent myotubule formation (Freed et al., 2005). BMSCs-exo treatment with CTF1 can also result in rapid STAT3 phosphorylation, leading to increased cellular adherence (Bortolotti et al., 2017). There is also prior evidence in ICR (Institute of Cancer Research) and B6 (C57BL/6) mice that CTF1 can promote STAT3 phosphorylation in the uterine luminal epithelium, leading to higher implantation rates (Kobayashi et al., 2014). These data highlight the importance of CTF1 as a promoter of angiogenesis and persistence following BMSCs-exo treatment in vitro and in vivo, thereby leading to improved endometrial regeneration and increased endometrial receptivity.