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Adipose extracellular matrix promotes skin wound healing by inducing the differentiation of adipose-derived stem cells into fibroblasts

Fibroblasts are the major effector cells of skin wound healing. Adipose-derived stem cells can differentiate into fibroblasts under certain conditions. In the present study, it was hypothesized that adipose-derived stem cells (ADSCs) could be induced by the adipose extracellular matrix (ECM) to differentiate into fibroblasts in order to promote skin wound healing. First, flow cytometry was used to detect the ratio of fibroblasts and relative expression of the fibroblast markers cytokeratin 19 (CK19) and vimentin in ADSCs. Then, the effect of the adipose ECM during the differentiation of ADSCs into fibroblasts was investigated by detecting the total amount of collagen fibers and degree of fibrosis, and the proliferation and cell cycle of differentiated fibroblasts, using the MTT assay and flow cytometry analysis respectively. Finally, a mouse skin wound model was established and treated with PBS, ADSC suspension or ECM + ADSCs to compare wound healing rate and expression of collagen I and collagen III by immunohistochemistry. Following induction of ADSCs with the adipose ECM, more fibroblasts were found, expression of CK19 and vimentin increased, and a greater degree of fibrosis occurred, which revealed the positive effect of the adipose ECM on the differentiation of ADSCs into fibroblasts. In addition, the induced fibroblasts had enhanced proliferation activity, with more cells in the S phase and fewer in the G2/M phase. The in vivo experiment indicated that the ECM produced by the ADSCs had a faster wound healing rate and increased expression of collagen I and collagen III compared with mice injected with PBS or ADSCs alone, which verified that ADSCs induced by the adipose ECM had a positive effect on skin wound healing. The present study demonstrated that the adipose ECM in combination with ADSCs may be a novel therapeutic target for the repair of skin injury, due to the ability of the adipose ECM to induce the differentiation of ADSCs into fibroblasts and to facilitate the wound healing process.


Introduction
Wound healing is a complex process in which tissue homeostasis and the protective role of the skin are restored (1). Skin is a complex organ that is composed of the epidermis, dermis, and skin appendages, while wound healing in adult mammals leads to scar formation without skin appendages (2). Wound healing is an extremely dynamic process that includes a variety of cellular and biochemical processes, of which dermis collagen remodeling and scar budding are two important parts of tissue repair during the maturation phase (3). As previously reported, several immune cells, including T cells, may be associated with scar formation, and epidermal cells have a critical role in wound healing by regulating the extracellular matrix (ECM) (4). Scars, which are harmful to normal tissue function, are created during skin wound healing and are specifically caused by excessive deposition of the ECM by fibroblasts and myofibroblasts (5). A previous study has noted that acute skin wound healing is a coordinated process and that keratinocytes and fibroblasts are closely related to epidermal and dermal repair (6).
Fibroblasts, as cells of mesenchymal origin, have an elongated morphology with numerous cell processes extending from their surface, and fibroblasts serve a pivotal role in the maintenance of ECM homeostasis under normal circumstances (7). It has been demonstrated that the heterogeneity of fibroblasts exerts positive effects on wound healing without scar formation and full recovery of the native tissue structures in fetuses and the oral mucosa (8). The critical role of fibroblasts in wound healing has been recognized through their generation of ECM and differentiation into myofibroblasts (9). Fibroblasts are a type of mesenchymal cell in connective tissue and have a significant role in depositing the collagen and elastic fibers of the ECM (10). Additionally, recent research has demonstrated that stem cell treatment can be used as a treatment approach for wound repair and tissue regeneration, such as adipose-derived stem cells (ADSCs) and bone marrow-derived stem cells, which have been studied under both pre-clinical and clinical conditions (11).
ADSCs are a class of cells that have multidirectional differentiation potential and are potential contributors to wound healing at both the cellular level in vitro and histological level in vivo; ADSCs are also easy to obtain from subcutaneous adipose tissues (12). A previous study has revealed that ADSCs have the capability to differentiate into a variety of cell types, and therefore, they are thought to be effective in regenerative medicine (13). Other research has demonstrated that ADSCs can be induced to differentiate into fibroblasts by regulating connective tissue growth factors in vitro (14). Therefore, the present study aimed to investigate the potential mechanism of adipose ECM induction of ADSC differentiation into fibroblasts, thereby promoting skin wound healing.

Materials and methods
Ethics statement. Informed consent was obtained from all the participants. The present study was approved by the Ethics Committee of General Hospital of Chinese PLA (Beijing, China; approval no. S2017-059-10) and conducted in accordance with the 2000 Helsinki Declaration. All experiments involving animals were approved by the Animal Ethics Committee of General Hospital of Chinese PLA (approval no. 2017-x13-25).
Study subjects. Adipose tissues were derived from five healthy adult women (age, 25-65 years; body mass index, <30; hemoglobin, >10 g/l) who were admitted for liposuction to the Department of Plastic and Reconstructive Surgery, General Hospital of Chinese PLA from January 2017 to March 2017. The cases had no diabetes mellitus, hypertension, serious systemic metabolic diseases or lipid disorders. Adipose tissues were selected from the abdomen and thigh. Patients were treated with tumescent anesthesia (tumescent solution: 1,000 ml of 0.9% normal saline + 10 ml of 2% lidocaine + 1 ml of 0.1% adrenaline). Fat was suctioned using liposuction needles (3 mm in diameter) and under negative pressure via a liposuction device. The obtained adipose tissues were stored in 50 ml centrifuge tubes under aseptic conditions at -80˚C after washing.
Preparation of human adipose ECM. All the digestion and extraction processes were conducted in a 37˚C and 120 r incubator, and 1% penicillin/streptomycin and phenylmethanesulfonyl fluoride were added to the acellular fluid. A total of 20 ml of obtained adipose tissues were placed in a 50 ml centrifuge tube and washed with 0.1 mmol/l PBS three times to remove blood and tumescent fluid. Then, the tissues were stored at -80˚C for 1 h and rewarmed at 37˚C for 40 min, with three freeze-thaw cycles. Next, the tissues were centrifuged at 7,000 x g for 3 min at room temperature, followed by the removal of the oil from the upper layer, lower liquid and precipitate. Then, the tissues were mixed with 0.25% trypsin/0.1% EDTA and stirred at 37˚C (120 r/min) overnight, followed by three washes in 0.1 mmol/l PBS, 30 min per wash. Next, 99.9% isopropanol was used for extraction for 36 h and was replaced every 12 h to remove lipids, followed by three washes in 0.1 mmol/l PBS, 30 min per wash. The tissues were detached with 0.25% trypsin/0.1% EDTA for 4 h and rewashed three times with 0.1 mmol/l PBS, 30 min per wash. Then, the tissues were treated with 1,000 U of nuclease overnight, followed by three washes with 0.1 mmol/l PBS, 30 min per wash, and 99.9% isopropanol was used for extraction for 6 h. Following three more washes with PBS, the obtained adipose ECM was placed on a plate and stored at -20˚C. A vacuum freeze-drying machine was used to dry the ECM for 24 h until further use. The ECM had a spherical, loose and porous structure. Parts of the ECM were powdered at 12,000 x g for 10 min and sterilized through anterior cobalt-60 γ-ray radiation.
Isolation and culture of human ADSCs (hADSC). hADSCs were selected from one patient and obtained using enzyme digestion. The steps were as follows: First, 0.1 M PBS was used to wash fat granule tissues to remove tumescent liquid, blood and cell debris; then, fat granules were cut and added to Low-glucose Dulbecco's Modified Eagle's Medium (L-DMEM; cat. no. 31600034; Beijing Unique Biotechnology Co., Ltd., Beijing, China) containing 0.1% type I collagenase and digested on a shaking table for 40 min at 37˚C (120 x g). L-DMEM at the same volume containing 10% fetal bovine serum (FBS; cat. no. 16000044; Thermo Fisher Scientific, Inc., Waltham, MA, USA) was utilized to terminate the digestion. Samples were allowed to separate into layers and filtered through a 200-mesh screen mesh. With the filtrate collected, centrifugation was performed at 1,500 x g for 10 min at room temperature and the supernatant was discarded, leaving precipitated cells behind. The cells were re-suspended in L-DMEM containing 10% FBS. After mixing and counting, the cells were inoculated on a 10 cm culture dish and cultured in an incubator at 37˚C with 5% CO 2 . After 24 h, the culture medium was replaced, and it was subsequently replaced every 2-3 days. When the hADSC confluence reached ~80%, a sub-culture was performed.
Induction and differentiation of hADSC. hADSCs from the third generation were selected and divided into two groups, with the cell density adjusted to 1x10 5 : Control group (cells inoculated and cultured in L-DMEM containing 10% FBS) and ECM group (cells inoculated in human adipose ECM material and cultured in L-DMEM containing 10% FBS). After culturing for 2 days, matrix material inoculated with ADSCs was transferred to a Transwell chamber under sterile conditions, and an appropriate amount of medium was added to the level of the microporous filter membrane to expose the matrix material on the gas-liquid level. Six parallel samples were used for each group. The medium was replaced every day. The survival condition of cells was observed after 1, 3, 5, 7 and 9 days under an inverted microscope, followed by subsequent studies.
Flow cytometry. The cultured ADSCs were treated with 0.25% trypsin. Then, the cell concentration was adjusted to 1x10 6 cells/ml. A total of 1 ml of cells was centrifuged at 403 x g for 10 min at room temperature, after which the supernatant was discarded and 2 ml of PBS was added. Centrifugation was performed again. After the supernatant was discarded, the cells were fixed with a pre-cooled ethanol solution with a volume fraction of 70% and incubated at 4˚C overnight. The next day, the cells were washed with PBS twice. Then, 100 µl of the cell suspension (no less than 10 6 cells/l L) was added along with 1 ml of propidium iodide dye at a concentration of 50 mg/l containing RNase. The cells were kept in the dark for 30 min and filtered with a 300-mesh nylon mesh. Cell cycle distribution was measured through detection of red fluorescence at an excitation wavelength of 488 nm using flow cytometry (6HT; Cellwar Bio-technology Co., Ltd., Wuhan, Hubei, China). The obtained data from flow cytometry were analyzed by applying the BD FACSDIVA V8.0.1 software (BD Biosciences, San Jose, CA, USA).
MTT assay. When the confluency of the ADSCs reached ~80%, the cells were washed with PBS twice and treated with 0.25% trypsin, followed by preparation of a single cell suspension. After counting, the cells were seeded in a 96-well plate with a cell density of 3-6x10 3 cells per well (0.2 ml, with 6 replicate wells). Then, the culture plate was removed after 24, 48 and 72 h of incubation, and the cells were incubated with another medium containing a 10% MTT solution (5 g/l; cat. no. GD-Y1317; Guduo Biotechnology Co. Ltd., Shanghai, China). After incubation for an additional 4 h, the supernatant was aspirated and 100 µl of DMSO (Sigma-Aldrich; Merck KGaA, Darmstadt, Germany) was added to each well with gentle shaking for 10 min to allow the formazan crystals produced by living cells to dissolve. The optical density (OD) values at 490 nm were measured using a microplate reader (Nanjing DeTech Experimental Equipment Co., Ltd., Nanjing, Jiangsu, China). Each experiment was repeated three times. A cell viability curve was plotted with the time points as the abscissa and OD values as the ordinate.
Reverse transcription-quantitative polymerase chain reaction (RT-qPCR). Total RNA was extracted using TRIzol (cat. no. 16096020; Thermo Fisher Scientific, Inc.), and then, according to the instructions of the RT-qPCR kit (cat. no. AB4105C; Verso 1-step RT-qPCR kit; SYBR-Green, ROX; Thermo Fisher Scientific, Inc.), 5 µg of RNA was reverse transcribed into cDNA. The target gene was amplified by PCR using a 25 µl reaction volume, including 300 ng of cDNA, 1X PCR buffer solution, 200 µmol/l dNTPs, 80 pmol/l forward primers, 80 pmol/l reverse primers, and 0.5 U of Taq enzyme (cat. no. S10118; Shanghai Yuanye Biological Technology Co., Ltd., Shanghai, China). The reaction conditions were as follows: Pre-denaturation at 94˚C for 5 min, 30 cycles of denaturation at 94˚C for 30 sec, annealing at 54.5˚C for 30 sec, and extension at 72˚C for 30 sec, followed by extension at 72˚C for 10 min and preserved at 4˚C. The primer sequences of collagen (Col) I, Col III and β-actin are listed in Table I. β-actin was used as the internal reference. Relative fold changes in mRNA expression were calculated using the formula 2 -ΔΔCq (15). The experiment was repeated three times.
Western blot analysis. Total protein was extracted from cells with a radio-immunoprecipitation assay buffer (cat. no. R0010; Beijing Solarbio Science & Technology Co., Ltd., Beijing, China) containing PMSF, incubated on ice for 30 min and centrifuged at 4˚C for 10 min (12,000 x g). The supernatant was obtained. The protein concentration of the supernatant of each sample was determined using a bicinchoninic acid kit (cat. no. 23225; Thermo Fisher Scientific, Inc.), and deionized water was then used to adjust the amount of protein. Next, a 10% SDS-PAGE gel (cat. no. P0012A; Beyotime Institute of Biotechnology, Shanghai, China) was prepared and 50 µg of the protein sample was added to each well. Electrophoresis was conducted at a constant voltage of 80 V for 2 h. The proteins were transferred onto polyvinylidene fluoride (PVDF) membrane (cat. no. ISEQ00010; EMD Millipore, Billerica, MA, USA) with a voltage of 110 V for 2 h. The PVDF membrane was blocked with TBST buffer containing 5% skimmed milk powder for 2 h. Then, the membrane was washed with TBST and incubated with rabbit polyclonal antibody Col I (1:100; cat. no. ab34710), rabbit polyclonal antibody Col III (1:100; cat. no. ab7778), and β-actin antibody (1:100; cat. no. ab8224; all Abcam) at 4˚C overnight, followed by three washed with TBST, 10 min each time. The membrane was then washed with 0.1% PBS/Tween-20 (PBST) 3 times at room temperature, 10 min each time, immersed in enhanced chemiluminescence reaction solution (cat. no. WBKLS0100; EMD Millipore), and developed. β-actin was used as the internal reference, and the ratio of the gray values of the target protein band to the internal reference band was used as the relative protein expression and analyzed by Quantity One v4.6.2 (Bio-Rad Laboratories, Inc., Hercules, CA, USA).
Immunofluorescence. The cells in each group were seeded into 24-well plates pre-coated with poly-L-lysine. After cell adherence, the medium was removed and the cells were gently washed with 0.01 mol/l preheated PBS three times. , on the back of each mouse, 2x1.5 cm full-thickness skin wounds were made. The 0.5 mm thick silicone resin sheet was cut into a concentric circle twice the size of the wound and fixed around the wound with nylon suture. In the preparation of the wound model, silica gel ring was used to fix the wound, so the interference of skin contraction to the experiment was limited to the maximum extent (16,17). Mice were randomly assigned into three groups (10 mice in each group): The PBS group (mice treated with an injection of 1 ml PBS at the wound base and wound edge), the Control group (mice treated with an injection of 1 ml ADSC suspension, 5x10 5 cells/ml) and ECM group (ECM powder, sterilized by cobalt-60 γ ray radiation, was immersed in the culture medium of ADSCs and balanced for 24 h in an incubator, and mice were evenly injected with 1 ml of ADSC-conditioned media with ECM at the base of the wound and wound edge). Mice with wounds were kept in individual cages to avoid biting each other. Mice were photographed 14 days following the injury to observe wound healing and these images were assessed visually only. The complete wound tissues were then removed, half of which was used to extract RNA for RT-qPCR and the other half were used for embedding, slicing, Masson staining and immunohistochemistry of Col I and III.
Immunohistochemistry. Wound tissue specimens from each group were fixed with 10% formalin for 5 h at room temperature, embedded in paraffin using conventional methods, sliced to a thickness of 3 µm and dehydrated with xylene and gradient alcohol. High-pressure antigen retrieval was conducted with citrate antigen repair solution (pH, 7.2-7.4). The slices were incubated at 4˚C overnight with primary rabbit polyclonal antibodies against Col I (1:100; cat. no. ab34710; Abcam) and Col III (1:100; cat. no. ab7778; Abcam) and then at 37˚C for another 1 h, followed by a subsequent rinse with PBS. Then, a biotin-labeled secondary antibody (1:200; cat. no. ab150077; Abcam) in working fluid was added and incubated at 37˚C for 30 min. Next, 3,3'-diaminobenzidine (DAB) was added for coloration for 10 min at room temperature, after which, the slices were stained with hematoxylin at room temperature for 2 min, dehydrated by gradient alcohol, mounted in neutral balsam, and observed under an optical microscope. The hysterosalpingography (HSG) positive reactant exhibited a tan color in the cytoplasm and nucleus of the cells. Immunohistochemically stained sections were observed under a light microscope (DMM-300D, Shanghai Caikon Optical Instrument Co., Ltd., Shanghai, China) (magnification, x400), and the mean OD values were calculated by Image Pro Plus pathological image analysis software (6.0), followed by statistical analysis (18). The experiment was repeated three times.
Masson staining. Adipose ECM material containing induced ADSCs at each time point and mouse skin wound specimens were fixed with 4% formalin at room temperature (cells for 30 min, tissues for 24 h), embedded in paraffin, and cut into 4 µm serial sections. Masson collagen fiber staining at room temperature for 3 min was used to observe the proliferation of collagen fibers in ECM materials and skin wound tissues. The differentiation effect in ADSCs induced by the ECM upon skin wound healing was analyzed through observation of collagen fibers under the optical microscope in x200 magnification power (Olympus Corporation, Tokyo, Japan). The four corners of the specimen and the percentage of collagen fibers in five central fields were calculated. The collagen fiber content was expressed as the percentage of collagen fibers in the view field, and the obtained mean value was used for statistical analysis.
Statistical analysis. SPSS 21.0 statistical software (IBM Corp. Armonk, NY, USA) was used to analyze the data, which were processed with normal distribution and homogeneity of variance tests. Measurement data are expressed as the mean ± standard deviation. Comparisons between two groups were conducted by t-test. Comparisons among multiple groups were assessed by one-way analysis of variance. The pairwise comparisons among groups were assessed by Tukey post hoc test. Enumeration data are presented as % and were assessed by the chi-square test. P<0.05 was considered to indicate a statistically significant difference.

Morphological observation and surface marker detection of
ADSCs. Initially, the morphology of ADSCs was observed by microscopy (Fig. 1A). Sparsely distributed single cells were found to be in adherent growth. Typical ADSCs exhibited fusiform, polygonal, spindle-shaped fibroblast-like appearance. The cell bodies were narrow and long, with a long spindle shape, were arranged orderly, and had a certain direction. They grew in a shoal shape. After 1 week of incubation, cells had adhered to the cell wall. Most cells were spindle-shaped fibroblast-like, some were triangular, and a few exhibited colony growth, while a few were in the round and oval cell growth phases. After 2 weeks of incubation, fibroblast-like cells dispersed and exhibited colony growth, and cells were arranged in a concentric circle, with whirlpool and radial shapes. The central cells of the colony were densely distributed, while they were loose around the periphery. At 4 weeks, the cell confluence reached ~80% (Fig. 1A). Flow cytometry was then employed to detect the distribution of ADSC-specific surface markers. The ADSCs expressed a wide variety of cell surface markers (Fig. 1B) (Fig. 1C).

ADSCs can differentiate into fibroblasts following induction with adipose ECM.
To investigate whether the ECM could induce the differentiation of ADSCs into fibroblasts, flow cytometry was used to detect the ratio of fibroblasts. Compared with the control group, the proportion of fibroblasts in the ECM group significantly increased, from 4.23% on day 3 to a peak of 16.43% on day 7 of induction, and decreased on day 9 ( Fig. 2A and B). Immunofluorescence staining revealed that CK19 and vimentin were mainly expressed in the cytoplasm. Compared with the control group, the relative expression of CK19 and vimentin was significantly increased in the ECM group (all P<0.05; Table II). In the ECM group, the expression of CK19 and vimentin reached a peak on day 7 post-induction and began to decline on day 9 ( Fig. 2C and D; Table II). The above results demonstrated that under the induction of the adipose ECM, ADSCs differentiated into fibroblasts.
Fibroblast proliferation is enhanced following induction with adipose ECM. To investigate the vitality of ADSCs that differentiated into fibroblasts following induction with the adipose ECM, MTT assays and flow cytometry analyses were performed to detect the proliferation activity and cell cycle distribution of the differentiated fibroblasts, respectively. As illustrated in the growth curve of Fig. 3A, the numbers of viable cells in the ECM and control groups were gradually enhanced over time, exhibiting a fast-growing period from 4-6 days post-induction, reaching a peak at 6-8 days, plateauing, and starting to decline from 14-16 days. There was no significant difference in the OD value between the control group and ECM group from 2-12 days post-induction (P>0.05). However, on days 14 and 16, compared with the control group, the ECM group displayed a significantly increased cell proliferation activity (P<0.05; Fig. 3A), suggesting that the ECM can maintain continuous cell proliferation activity.
Cell cycle phase distribution analysis was performed at the 9th day post-induction, and the results revealed that: Compared with the control group, the ECM group had a significantly reduced number of ADSCs in the early stage of synthesis Table II. Relative expression of CK19 and vimentin in ADSCs following adipose ECM induction, as determined by immunofluorescence staining.   (G 0 /G 1 phase) and in the late stage of DNA synthesis (G 2 /M phase), while an increased number of cells were arrested in the S phase (P<0.05; Fig. 3B and C). The above results suggested that fibroblast proliferation was enhanced following induction with the adipose ECM, since most of the cells were in the synthesis phase (S phase).

ECM induction upregulates collagen expression in ADSCs.
Then, to determine the effects of the adipose ECM induction of ADSCs on wound healing in vitro, RT-qPCR and western blot analyses were conducted to determine the expression of the wound healing-related factors Col I and Col III in induced ADSCs. Compared with the control group, the mRNA and protein expression of Col I and Col III in the ECM group was significantly increased at 12 days post-induction (P<0.05; Fig. 4A-C). The expression of Col I and Col III was obviously upregulated in ADSCs that differentiated into fibroblasts following induction with adipose ECM.

ECM-induced ADSCs promote wound healing in vivo.
Lastly, a mouse skin wound model was established to further explore the effect of ADSCs on skin wound healing. Wound healing in each group was observed by the naked eye (Fig. 5A). No significant change was observed between the PBS and control groups; compared with the PBS and control groups, the wound area of the ECM group was significantly reduced. RT-qPCR and immunohistochemistry were performed to determine the expression of Col I and Col III. Compared with the PBS and control groups, positive expression of Col I and Col III in the ECM group was significantly higher both at the mRNA and protein level (Fig. 5B, C and E). Masson staining was applied to detect the degree of fibrosis in mouse wound tissues, in order to explore the role of induced ADSCs in skin wound healing. The ECM group exhibited a notably elevated fibrosis level in wound tissues compared with the PBS and control groups ( Fig. 5D and F). The above results suggested that the expression of Col I and Col III protein and the fibrosis level  in wound tissues were upregulated following induction with adipose ECM, and subsequently the healing of the wounded skin was promoted.

Discussion
In the present study, the results demonstrated that induction of ADSCs with adipose ECM increased the expression of CK19 and vimentin, the proliferation activity, the expression of Col I and Col III, and enhanced skin wound healing in vivo, by inducing ADSCs to differentiate into fibroblasts. Skin wound healing is a dynamic and complicated process that involves the coordination of various cell types and cellular processes (19). ADSCs, multi-potent stem cells similar to bone marrow-derived mesenchymal stem cells, are easy to isolate and are efficient in promoting skin wound healing (20). Recent evidence has demonstrated that ADSCs are essential in the repair and regeneration of numerous damaged tissues, including the regeneration of skin tissue, by affecting fibroblasts (21).
Initially, the data obtained during the present study demonstrated that the adipose ECM effectively induced ADSCs to differentiate into fibroblasts. The results revealed that the proportion of fibroblasts and expression of their surface marker proteins CK19 and vimentin increased significantly after ADSCs were induced by the adipose ECM. A previous study has noted that although adult stem cells have a more limited differentiation potential than embryonic stem cells, they have the capability to differentiate into cells of different germ layers under appropriate stimulation (22). Human adult mesenchymal stem cells are rich in adipose tissue, which can be obtained by liposuction, and ADSCs can differentiate into different mesenchymal lineages, including cartilage, bone, muscle, and fat (23). Another study has also mentioned that ADSCs can be differentiated into various cell lineages, for instance, adipocytes, osteoblasts, chondrocytes, and myocytes, indicating the significant role of ADSCs in regeneration medicine and tissue engineering (24). In addition, ADSCs are regarded as multipotent stem cells and have great potential for the treatment of numerous serious human diseases, such as spinal cord injury and stroke (25). ADSCs have some features similar to mesenchymal stem cells from bone marrow and cord blood in terms of their multipotency and differentiation into endothelial cells and fibroblasts (26). Furthermore, increasing evidence has demonstrated that the differentiation of ADSCs is closely related to microenvironmental factors, including soluble and matrix-bound growth factors, the ECM, as well as neighboring cells (27). Recent data have demonstrated that when stem cells are exposed to the intrinsic properties of the ECM, they can differentiate into cells in mature tissues, and the ECM parameters are reported to be highly dynamic during their development, so that they exert a great influence on regulating the differentiation and arrangement of cells (28).
The present study also proposed that the differentiation of ADSCs into fibroblasts induced by adipose ECM promoted skin wound healing. Upregulated expression of the wound healing factors Col I and Col III was observed in differentiated fibroblasts, and the proliferation activity of fibroblasts and collagen fibers was also increased. The ECM of specific tissues induces cell proliferation, regulates the function of cells and drives cellular differentiation (29). It has been reported that hADSCs, which are abundant in adipose tissue, function as a source of cell treatment in terms of regenerating damaged tissues due to their ability to self-renew and differentiate into different cell lineages (30). Other authors have reported that human ADSCs are able to stimulate the proliferation and migration of dermal fibroblasts and enhance their production of matrix factors and thus accelerate the re-epithelialization of wounds in nude mice (21). A previous study has assessed the differentiation capability of ADSCs and their potential for wound healing in vivo (31). As previously discussed, ADSCs can differentiate into myofibroblasts and then re-differentiate into fibroblast-like cells under the regulation of growth factors, indicating the role of ADSCs in reducing the risk of fibrosis when finding new treatment strategies (32). In addition, collagen, as the basic protein used to form the connective tissues matrix, is capable of promoting the migration of keratinocytes to benefit the rebuilding of the damaged epidermis, and collagen also serves a pivotal role in accelerating wound healing (33). Mounting evidence indicates that collagens mediate many changes in the ECM during wound healing to facilitate tissue regeneration (34). Synthesis of the ECM is a key step in wound healing and is required to maintain the suppleness and mechanical properties of the skin and its major components, including collagens, while fibroblasts are involved in collagen production (35). Numerous data have demonstrated that skin wound healing is closely associated with matrix synthesis and degradation through the regulation of dermal fibroblast activity, and fibroblasts can synthesize many ECM proteins, such as collagen, and remodeling enzymes as well as their inhibitors (36). It has also been elucidated that dermal fibroblasts have a critical part in wound healing by accurately regulating their function (37).
In conclusion, the adipose ECM upregulates the expression of Col I and Col III in ADSCs and accelerates skin wound healing by inducing the differentiation of ADSCs into fibroblasts. Therefore, the present study provides a potential novel therapeutic target for skin tissue regeneration and skin wound healing. However, further studies are required to fully understand the specific mechanisms of ADSC differentiation for the enhancement of skin wound healing.