Lyophilized Catechol–Chitosan Mucoadhesive Hydrogels Loaded with Dental Follicle-Derived Mesenchymal Stem Cells Enhance Regenerative Healing of Palatal Donor Wounds

TL;DR

A new hydrogel containing dental follicle stem cells improves healing of palatal wounds in rats.

Contribution

A ready-to-use lyophilized hydrogel loaded with dental follicle-derived mesenchymal stem cells is developed for palatal wound healing.

Findings

DFMSC-loaded hydrogels showed over 80% cell viability after rehydration.

Palatal wounds healed faster with reduced inflammation and increased epithelial regeneration.

Cell viability remained above 70% even after five months of storage.

Abstract

Mesenchymal stem cells (MSCs) are candidates for the treatment of palatal wounds in combination with biomaterials. In this study, we developed a method for the production of a ready-to-use mucoadhesive hydrogel containing MSCs for palatal wounds and evaluated its healing effects. Dental follicle MSCs (DFMSCs) were isolated from the dental follicle tissue of a healthy twenty-year-old donor. DFMSCs were suspended in a cell-preserving solution containing platelet-rich plasma, trehalose, and DMSO, and loaded into a catechol–chitosan hydrogel solution at a ratio of 1:400 (v/v) with 5 × 105 or 6 × 106 cells per hydrogel to create a novel lyophilization method for cell integration into the biomaterial. Hydrogels were fabricated as scaffolds with a diameter of 5 mm and a depth of 4 mm. After lyophilization of the hydrogels with cells, a viability test was performed after the production of hydrogels on the seventh day and the fifth month. Palatal wounds were created as full-thickness wounds in rats using a 5 mm diameter punch. The hydrogels were applied to the palatal wounds of rats, and histochemical and immunohistochemical analyses were performed. The results showed that, after rehydration of the hydrogels, DFMSCs had over 80% viability and were homogeneously distributed in the hydrogels. After the application of DFMSC-loaded hydrogels, palatal wounds healed within 7 days, and inflammatory cell infiltration, fibroblastic proliferation, and ulceration were significantly reduced, while epithelial regeneration was significantly increased compared to the control group. The viability ratio of DFMSCs was 83.7% on the seventh day and 71.3% in the fifth month. Hydrogels loaded with DFMSCs represent a promising, ready-to-use biomaterial approach for supporting palatal wound healing.

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1. Introduction

Gingival recession is characterized by the positioning of the gingival margin apical to the cementoenamel junction and can lead to clinical problems such as esthetic issues, dentin sensitivity, and root surface caries [1]. Autogenous soft tissue grafts, commonly used in the treatment of gingival recession, are primarily harvested from the palatal region. However, after harvesting these grafts, the palatal region becomes a wound environment carrying morbidity factors such as pain, bleeding, necrosis, sensory disturbance, and risk of infection [2,3]. These complications not only reduce patient comfort but also prolong the healing process and can negatively impact clinical outcomes [3,4]. Significant innovations in surgical techniques and graft materials have been developed in recent years to reduce these complications. Single-incision and de-epithelialized single-incision techniques, defined as alternatives to the classical trap-door method, have significantly reduced postoperative pain and morbidity by shortening the operative time and reducing tissue trauma [4]. In addition, ensuring hemostasis in the donor area before surgery with palatal pre-suture applications facilitates bleeding control and accelerates healing [3].

Currently, the use of tissue-engineered biomaterials offers a promising approach to reducing the need for autogenous grafts. Collagen matrices, acellular dermal matrices, and other biological substitutes can provide clinically adequate soft tissue thickness and root surface coverage while eliminating donor site morbidity [5,6]. Additionally, the application of platelet-rich fibrin or growth factor-enriched regenerative materials has the potential to accelerate epithelialization and reduce postoperative pain and edema [7]. However, although autogenous grafts remain the gold standard for soft tissue regeneration, biologically supportive approaches are being investigated to reduce donor site morbidity and accelerate healing [8].

Wound healing involves complex, overlapping phases such as hemostasis, inflammation, proliferation, and tissue remodeling [9]. This process requires the synchronization of events such as new tissue formation, angiogenesis, cell migration, and matrix synthesis [9,10]. The role of mesenchymal stem cells (MSCs) in wound healing is also evaluated in terms of their capacity to support these phases [11,12]. These cells have been shown to be effective in regulating the immune response, suppressing inflammatory cytokines, and increasing anti-inflammatory factors. Furthermore, it has been reported that angiogenesis is stimulated and fibroblast proliferation and migration are supported through the secretion of paracrine mediators such as vascular endothelial growth factor and angiopoietins [13,14]. It is emphasized that MSC-derived exosomes can have significant effects on immunomodulation and angiogenesis in chronic wounds, and that this effect can be optimized with engineering approaches [15]. Owing to the immunomodulatory capacity, high proliferative potential, and differentiation ability of dental follicle-derived mesenchymal stem cells (DFMSCs), they can create paracrine effects that accelerate epithelialization and support angiogenesis, and these features make them a promising cellular source in oral tissue engineering and wound healing models [16,17].

Biomaterials play a critical role as drug delivery systems to ensure better adhesion of MSCs to the wound site, their viability, and their ability to maintain paracrine functions. Hydrogels stand out in this context [18]. MSC-loaded hydrogels have been shown to provide local release control by increasing cell retention and contributing to the regulation of proliferation, cell migration, angiogenesis, and inflammation during wound healing [19]. For this purpose, the combination of chitosan, known for its biocompatibility and biodegradability, and catechol groups, which provide strong tissue adhesion, is frequently preferred in the development of mucoadhesive and cell-friendly hydrogel systems for the wound site [20,21].

However, technical limitations related to long-term storage and formulation stability pose a significant obstacle to the translation of MSC-containing hydrogel systems into clinical use. Therefore, technological improvements to maintain shelf life and structural stability while supporting the biological performance of biomaterial systems have been focused on [22,23]. In this context, lyophilization is a method that reduces the logistical difficulties arising from the freeze–thaw process by providing hydrogel systems with long shelf lives and ready-to-use forms [24]. In studies investigating the functional properties of lyophilized hydrogels, hydrogels lyophilized with disaccharides have demonstrated improved stability and release properties, and could be used as successful drug delivery systems [25,26]. It is also known that drug loading into lyophilized hydrogels is easier and more efficient, and more importantly, the swelling ratio and controlled drug release potential are preserved [27].

The literature demonstrates that biological approaches to accelerate wound healing at palatal donor sites are limited, and drug delivery systems containing lyophilized DFMSCs have not been investigated for this purpose. Existing studies have focused primarily on bone or cutaneous tissue models, and data on oral mucosa-specific cellular hydrogel applications are limited. Therefore, we aimed to develop a DFMSC-loaded lyophilized mucoadhesive catechol–chitosan hydrogel system to promote palatal donor site healing in an in vivo rat palatal wound model and to evaluate its biological efficacy in vivo. We hypothesize that this cellular hydrogel will provide faster epithelialization, balanced inflammation, and increased angiogenesis compared to control groups.

2. Materials and Methods

2.1. Dental Follicle Mesenchymal Stem Cell Isolation

Dental follicle tissues were obtained from systemically healthy 20-year-old donors with no history of genetic or autoimmune diseases undergoing extraction of fully impacted third molars. The study protocol was approved by the Medical Sciences Ethics Committee of Muğla Sıtkı Koçman University with approval number 113 dated 27 August 2024. DFMSCs were isolated as described in our previous study [28]. In brief, dental follicle tissue was cut into 0.5–1 mm pieces and enzymatically digested using 3 mg/mL collagenase in a phosphate-buffered solution (PBS) for 45–60 min at 37 °C. After incubation, complete DMEM (DMEM low glucose supplemented with 10% fetal bovine serum and 1% Penicillin/Streptomycin 10,000 U) was added to inactivate the enzymatic reaction. A cell pellet was obtained by centrifugation at 1200 rpm for 5 min. The cell pellet was rinsed twice with complete DMEM and then cultured with complete DMEM in a T25 flask until the cells reached 80% confluence. Each passage was performed by trypsinizing cells with 0.25% Trypsin EDTA. The cells at the third passage were used for all the encapsulation procedures and animal experiments.

2.2. Characterization and Multilineage Differentiation Analyses

Flow cytometric characterization of DFMSCs was performed in accordance with the minimal criteria for MSCs defined by the International Society for Cellular Therapy (ISCT). Analyses were performed using a flow cytometer for cells at the third passage, after the cells had been removed from the bottom of the culture dishes with 0.25% trypsin EDTA. Fluorescein-conjugated monoclonal antibodies specific for designated cell surface markers, along with 10 μL of appropriate isotype controls, were added and incubated at room temperature (in the dark) for 15 min. After incubation, PBS was added and the mixture was centrifuged. The cell pellet was resuspended in 500 μL of washing solution. Flow cytometer analysis was performed using Accuri C6 plus Software. CD29 (PE), CD105 (FITC), CD90 (PerCp), and CD73 (APC) antibodies were used as positive markers for MSCs. Additionally, CD14 (PE), CD34 (APC), CD45 (PerCp), and HLA-DR (FITC) were used as negative markers. All antibodies were obtained from BD Biosciences, San Jose, CA, USA.

The osteogenic differentiation protocol was applied as follows: Third-passage cells were incubated with complete DMEM in 6-well plates at a density of 200,000 cells/well until reaching 80–90% confluence. Then, the medium was removed and 2 mL of Osteogenic Differentiation Medium (StemPro Osteogenic Differentiation Kit, Thermo Scientific, Waltham, MA, USA) was added to each well. The cells were incubated at 37 °C in a 5% CO_2_ incubator according to the kit manufacturer’s protocol for 21 days. The cells were stained with Alizarin Red to observe calcium deposits. The chondrogenic differentiation protocol was as follows: A 2 mL volume of Chondrogenic Differentiation Medium was added to each well. Incubation was carried out at 37 °C in a 5% CO_2_ incubator. The medium was changed every 3 days, and incubation continued for 21 days. The formation of cartilage matrix components was observed using the Alcian Blue staining protocol. The adipogenic differentiation protocol was as follows: A 2 mL volume of Adipogenic Differentiation Medium was added to each well. Incubation was carried out at 37 °C in a 5% CO_2_ humidified chamber. The medium was changed every 3 days, and incubation continued for 14–21 days. At the end of the culture period, the formation of adipocytes and oil droplets was observed using the Oil Red staining protocol.

2.3. Fluorescence Cell Labeling

To track the DFMSCs in vivo and analyze their distribution in the hydrogels, the cells were labeled with a Qdot marker (Qtracker^®^ 655 Cell Labeling Kit, Thermo Scientific, Waltham, MA, USA), which results in fluorescent nanocrystals in the cytoplasm. The Qdot labeling of MSCs was performed according to the kit manufacturer’s protocol. Cells were suspended in 10 mL of medium (DMEM, 10% FBS, 1% penicillin/streptomycin) and 5 nmol/L Qdot per 1 × 10^6^ cells and incubated for 1 h at 37 °C in a 5% CO_2_ chamber. Qdot-labeled cells were cultured for 24 h, labeled with DAPI, and analyzed under a fluorescence microscope.

2.4. Platelet-Rich Plasma Isolation and Quality Tests

Cardiac blood samples were collected from rats to isolate platelet-rich plasma (PRP) before the production of the hydrogel, which contained or did not contain MSCs, for use in an experimental palate defect animal model. The lyophilization solution for the cells contained 60% PRP, 35% Trehalose, and 5% DMSO. The PRP isolation protocol was performed as described previously [29]. Briefly, 6–10 mL cardiac blood samples were collected from rats in a tube containing 3.2% sodium citrate. The blood sample was centrifuged at 1500 rpm for 10 min at +4 °C, and the plasma and buffy coat were collected with a pipette. The collected sample was transferred to a sterile 15 mL Falcon tube and centrifuged at 2000 rpm for 10 min at +4 °C. After centrifugation, the upper 3/4 of the tube was discarded, and the bottom 1/4 was collected to obtain PRP. For quality tests, 50 microliters were separated from each sample of the obtained PRP, and the following tests were performed: (1) platelet count; (2) growth factor analysis. The platelet count was determined using a platelet counter (PC100 Platelet Counter, Dutch Medical Devices BV, Valkenswaard, The Netherlands), and growth factor analysis was performed using an ELISA kits for PDGF (Abcam, Cat# ab267585, Cambridge, UK) and VEGF (Abcam, Cat# ab100786, Cambridge, UK).

2.5. Hydrogel Production and Characterization Analyses

Catechol grafting: For hydrogel production, firstly, catechol-functionalized chitosan (catechol–chitosan), whose chemical structure is given in Supplementary Figure S1, was synthesized. The catechol functionalization of chitosan was achieved via carbodiimide-mediated coupling chemistry. Briefly, hydrocaffeic acid was conjugated to the primary amine groups of chitosan using N-(3-dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride (EDC) as a coupling agent, enabling amide bond formation between the carboxyl groups of hydrocaffeic acid and the chitosan backbone. The reaction was conducted under mildly acidic conditions (pH ~5.5) to favor carbodiimide activation and preserve catechol functionality. For this purpose, 60 mL of 1% (w/v) chitosan (medium molecular weight, 85% deacetylation, Sigma, Saint Louis, MO, USA) solution was prepared in 1% acetic acid and magnetically stirred for 72 h until complete homogenization. For catechol grafting, 0.145 g of hydrocaffeic acid (>98%, Sigma) and 0.356 g of N-(3-dimethylaminopropyl)-N’-ethylcarbodiimide hydrochloride (>98%, Sigma) were dissolved in 15 mL of ethanol and 15 mL of deionized water, respectively. These two solutions were mixed and added to the chitosan solution under vigorous stirring. The pH of the mixture was adjusted to 5.5 using 1 M NaOH. The reaction was continued for 18 h, during which the initially transparent solution turned light brown, consistent with catechol grafting as previously reported [30]. Following the coupling reaction, the catechol–chitosan conjugate was purified to remove unreacted hydrocaffeic acid, residual coupling agents, and low-molecular-weight byproducts. The reaction mixture was first diluted with an acidic aqueous solution (pH 5.0) to stabilize the catechol groups and prevent oxidation. The solution was then subjected to dialysis using a regenerated cellulose membrane (molecular weight cut-off: 12–14 kDa) against acidic distilled water (pH 5.0) for 72 h, with frequent medium replacement. The purified product was then lyophilized, yielding a fibrillar-layered morphology distinct from that of native chitosan. The synthesized catechol–chitosan was stored at −20 °C until use. To confirm catechol–chitosan modification, the prepared catechol–chitosan solution was first analyzed by UV-Vis (Thermoscientific-MultiScan-GO, Waltham, MA, USA) in the wavelength range of 200–1000 nm. The UV-Vis spectrum of the catechol-modified chitosan exhibited a characteristic absorption peak at approximately 280 nm, which can be attributed to π → π* electronic transitions of the aromatic catechol moiety originating from hydroxycaffeic acid conjugation (Figure 1A). This absorption band indicated the successful grafting of catechol functional groups onto the chitosan backbone. The appearance of this peak is consistent with previous studies reporting catechol–chitosan conjugation [30].

DFMSC-loaded hydrogel production and lyophilization: Hydrogel production was initiated by preparing a 2% (w/v) catechol–chitosan solution in 1% acetic acid, and the solution was magnetically stirred for 48 h until complete homogenization. A 20 mg/mL genipin (≥98.0%, Glentham Life Sciences, Corsham, UK) solution was prepared in ethanol as a crosslinking agent. For the preparation of cell-loaded hydrogels, cells were first incorporated into the genipin solution prior to crosslinking. The solutions were first passed through a 0.22 μm filter and sterilized by exposure to ultraviolet light for 30 min under aseptic conditions before the cells were loaded into the hydrogels. Cells were prepared as 500,000 DFMSCs or 1 million DFMSCs containing DMSO + PRP + trehalose (60% PRP, 35% Trehalose, and 5% DMSO)/hydrogel (1:10 v/v) to be loaded into the prepared hydrogels at 200 μL. To investigate the effect of DFMSCs, hydrogel carriers were produced both with and without DFMSC loading, and the difference between their effects was taken into account.

The resulting mixture was then transferred to 96-well microplates in 200 µL volumes using a micropipette and used as templates to obtain hydrogels of the same size. To prepare for the lyophilization process, the hydrogels were first frozen at −80 °C for 12 h. The frozen contents were dried using a lyophilizer (Yamato DC 801, Freeze Dryer, Yamato Scientific, Tokyo, Japan) at −100 °C and 4 Pascal pressure for 24 h [31].

Hydrogel characterization and Biocompatibility tests: Within the scope of hydrogel characterization studies, physical characterization, swelling and degradation tests, mucoadhesion tests, in vitro DFMSC-release tests, in vitro cytotoxicity tests, and biocompatibility tests were performed.

FTIR Analysis: FTIR analysis of the lyophilized hydrogels was performed using a Perkin-Elmer Spectrum 65-FTIR spectrometer (Shelton, CT, USA) in the 400–4000 cm^−1^ range. FTIR analyses were performed separately for the prepared unloaded catechol–chitosan hydrogels and the DFMSC-loaded hydrogels, and the functional group absorption bands of the hydrogels and possible functional group changes were evaluated by comparing the obtained spectra.

SEM Analysis and Image Processing-Based Particle Counting: Morphology and surface topography studies of unloaded and DFMSC-loaded hydrogels were performed with a JSM-7600F Scanning Electron Microscope (SEM) (JOEL, Tokyo, Japan). To quantify these observations, image processing-based particle counting was performed using ImageJ version 1.53t (National Institutes of Health, Bethesda, MD, USA), Python version 3.9.0, and Matplotlib version 3.5.1 heatmap software. Briefly, the image was converted to grayscale, and after blurring, adaptive thresholding was used to separate particles from the background. Each particle was counted individually using a contour detection algorithm. The image was divided into a 10 × 10 grid (100 fields), and the number of particles in each cell was determined.

Swelling and Degradation Tests: The swelling and degradation behavior of the hydrogels were evaluated over time at 37 °C and pH 6.8, reflecting the physiological conditions of the oral environment, in accordance with previously reported salivary pH values [32]. For the swelling tests, the initial dry weights (W_initial_) of the lyophilized hydrogels, both DFMSC-loaded and unloaded, were recorded. Each hydrogel was then placed in 5.0 mL of PBS in sealed containers and incubated at a constant temperature. At predetermined time intervals, the hydrogels were removed from the swelling medium using an aluminum mesh, excess surface liquid was removed with absorbent paper, and wet weights (W_wet_) were recorded for 12 h. Measurement was performed at 24 h to determine the equilibrium swelling ratio. The % swelling behavior of the hydrogels was calculated over time using the following equation:

$$[eqn]$$

The time-dependent degradation behavior of the hydrogels was assessed in parallel with the swelling tests. Initially, the dry weights (Winitial) of DFMSC-loaded and unloaded lyophilized hydrogels were recorded, and hydrogels were placed in 5.0 mL of PBS in sealed containers and incubated at constant temperature. Degradation tests were continued for 12 h, during which the hydrogels were periodically removed from the medium with an aluminum mesh at predetermined time intervals, excess surface water was removed with absorbent paper, and the gel was subsequently re-dried. Then, the dry weights (Wfinal) of the hydrogels were recorded. An additional measurement was conducted at 24 h to evaluate the extended degradation behavior. The percentage degradation was calculated as a function of time using the following equation:

$$[eqn]$$

Mucoadhesion test: Commercially available calf buccal tissue was used to evaluate the mucoadhesive performance of the hydrogels and the influence of catechol modification on its adhesion properties. The buccal tissues were first mounted onto a supporting plate. Hydrogels produced with catechol–chitosan and unmodified chitosan were then manually pressed onto the tissue surface to ensure initial adhesion in separate sets, eight from each group [33]. The tissue–hydrogel assemblies were placed into a beaker, which was filled with pH 6.8 PBS pre-equilibrated to 37 °C until the samples were fully submerged (Supplementary Figure S2). Magnetic stirring was performed at 200 rpm throughout the test period, and the number of hydrogels remaining adhered to the mucosa was recorded at 10 min intervals [34].

In vitro cumulative MSC release test: An initial calibration study was conducted to quantify the MSCs released using the absorbance data of MSCs released from the hydrogels. For this purpose, 18 standard solutions of MSC-PBS were prepared with concentrations ranging from 1 million cells/mL to 0.03 million cells/mL through serial dilution. Each standard solution was analyzed using a Thermoscientific-MultiScan-GO UV-Vis spectrophotometer over 200–1000 nm wavelengths. The analysis revealed a characteristic absorbance peak for MSCs at 278 nm. Therefore, 278 nm was selected as the analytical wavelength for the tests. The in vitro cumulative DFMSC release kinetics were evaluated separately for hydrogel groups loaded with 500,000 DFMSCs (1 × DFMSC) or 1 million DFMSCs (2 × DFMSC) using the DMSO + PRP + Trehalose formulation by using pH 6.8 PBS at 37 °C as a release medium. Approximately 10 mg of each hydrogel sample was immersed in 20 mL of PBS in sealed containers and incubated under agitation at 200 cpm. At predetermined time intervals (every 10 min during the first hour, every 30 min up to 6 h, and every 6 h thereafter), 4 mL aliquots were withdrawn for analysis. The absorbance of the collected samples was measured at 278 nm. To eliminate potential spectral overlap between hydrogel components and DFMSCs, the absorbance values of unloaded hydrogel samples were used as blanks. The absorbance values obtained at each time point were converted to DFMSC concentrations using the previously derived calibration curve.

In vitro Biocompatibility test: This test was performed based on the study of Schossleitner et al. [35]. The human keratinocyte cell line in our cell isolates was suspended in keratinocyte growth medium, seeded at 1 × 10^5^ cells/well in 24-well culture plates, and cultured until confluent for 24 h. After 24 h, a 1 × 1 cm^2^ lyophilized hydrogel or a lyophilized hydrogel containing lyophilization solution was placed on the confluent cells in replicates of 5 and cultured with the culture medium for 24 h, 7 days, and 14 days. At the end of the culture period, the cells were stained with CalceinAM (green) for live cells and Ethidium Homodimer-1 “EthD-1” (red) for apoptotic cells. Images were taken under a fluorescence microscope [35].

MTT Cytotoxicity Test: This test was performed based on the study of Lim et al. [36]. The 5 mm diameter lyophilized hydrogel was washed three times with a phosphate-buffered solution containing 1% penicillin/streptomycin and exposed to ultraviolet light for 2 h. Human skin fibroblasts (HDFs) were used for in vitro culture. A total of 10 × 10^3^ cells were seeded into 96-well culture plates, and the HDFs were cultured in Dulbecco’s Modified Eagle Medium (DMEM) containing 10% FBS and 1% Penicillin/Streptomycin in a CO_2_ incubator (37 °C, 5% CO_2_). When the HDFs reached 80% confluence, they were trypsinized with a 0.25% trypsin EDTA solution. For cell cytotoxicity tests, 5 mm (diameter) lyophilized hydrogels were placed in 96-well plates and cultured with 10 × 10^3^ HDF cells for 3 days. At the end of the culture period, the DMEM was discarded, freshly prepared DMEM was added, and new DMEM with MTT solution (100 μL/well, 0.5 mg/mL) was added to each well and incubated for 4 h. The medium was discarded, and 100 μL of DMSO was added to dissolve formazan. After all the formazan crystals had dissolved, 100 μL aliquots were transferred to a 96-well culture plate with five replicates per sample. The absorbance was measured at a wavelength of 570 nm using a microplate spectrophotometer [36].

Analysis of the distribution of DFMSCs within the hydrogel: The homogeneous distribution of the DFMSCs within the hydrogel was analyzed using confocal microscopy. Because DFMSCs were labeled with a quantum dot fluorescent marker before being added to the hydrogels, fluorescence was obtained at a wavelength of 594 nm during imaging. To visualize the cells within the lyophilized hydrogel with a confocal microscope, frozen-sectioning was performed, followed by confocal microscopy analysis.

Viability analysis of DFMSCs loaded into the hydrogel: We performed flow cytometry analysis for the viability of DFMSCs 7 days after the lyophilization of the hydrogels. Briefly, 4 mm × 5 mm hydrogels containing 5 × 10^5^ DFMSCs were rehydrated with 1 mL of DMEM containing 20% FBS and incubated for 12 h at 37 °C in a humidified 5% CO_2_ chamber with a rotator (200 cpm). After the incubation period, the released cells were centrifuged at 1500 rpm for 5 min. The remaining cell pellet was stained with Annexin V-7AAD antibodies (BD Biosciences, #556422) for 15 min in the dark at room temperature, as described in the manufacturer’s protocol. The stained cells were analyzed by flow cytometry (BD Biosciences, San Jose, CA, USA, Accuri C6 Plus). The lower left quadrant was analyzed for viable cells. Data are presented as a percentage (%) of cells.

2.6. Experimental Palatal Wound Model

The experimental animal procedures were approved by the Local Ethics Committee for Animal Experiments of the Experimental Animals Application and Research Center of Muğla Sıtkı Koçman University, with approval number 16/24, dated 8 August 2024. The number of experimental animals was determined using G-power sample size analysis to obtain statistical data based on these variables and the 3R rule. In this study, two sacrifice times were determined based on palatal/mucosal wound healing times: days 7 and 14. Accordingly, the experimental study consisted of six groups and two separate sacrifice times. The effect size was set at 0.50, the type I error at 0.05, and the power at 0.80, as recommended for clinical studies, and the total number of animals was calculated as 60. The number of animals per group was 10 rats. Five of the rats per group were sacrificed on day 7, and five of the rats per group were sacrificed on day 14. The groups were as follows: Group 1: Palatal Wound; Group 2: Palatal Wound + Lyophilized Hydrogel; Group 3: Palatal Wound + Lyophilized Hydrogel with Cell Lyophilization Solution (PRP + Trehalose + DMSO); Group 4: Palatal Wound + Lyophilized DFMSCs; Group 5: Palatal Wound + Lyophilized Hydrogel with 500.000 DFMSCs; Group 6: Palatal Wound + Lyophilized Hydrogel with 1 million DFMSCs. The experimental palatal wound model was constructed based on a previous study [37]. For the experimental model, 3-month-old (average weight of 200 g) Wistar albino male rats were anesthetized with the intraperitoneal administration of 10 mg/kg xylazine hydrochloride (Rompun, Bayer 23.32 mg/mL), followed 10 min later by the intraperitoneal administration of 70 mg/kg ketamine hydrochloride (Ketalar, Parke-Davis, Detroit, MI, USA, 50 mg/mL). The animals were placed in the Trendelenburg position to prevent blood from the wound model from escaping into the trachea. The mouth was opened, and a 5 mm punch was used to create a secondary wound in the palatal mucosa in the middle part of the palate, removing the mucoperiosteal tissue. A standardized full-thickness mucoperiosteal defect with a diameter of 5 mm was created in the mid-palatal region, posterior to the maxillary incisors, using a biopsy punch. The defect involved the complete removal of the palatal mucoperiosteum, resulting in the exposure of the underlying palatal bone.

Wound Contraction: On days 7 and 14 of the experimental study, the wound closure area/wound area of the rats’ palates was measured macroscopically with a ruler and photographed for wound contraction. Wound contraction was calculated with the following formula: Wound area percentage (day X) = wound area mm^2^ on day X/wound area mm^2^ on day 0 × 100.

2.7. Histopathology and Immunohistochemistry

Palate tissues were taken from sacrificed rats and preserved in 10% formaldehyde. After the tissues were processed through an alcohol series, 4–5 µm sections were taken after paraffin blocking. Hematoxylin–eosin staining was performed to examine the tissue’s integrity and histology [38]. Microscopic images of epithelial integrity, fibroblastic proliferation, vascularization, inflammatory cell accumulation, and ulceration were evaluated. The scores were as follows: Ulceration and epithelial integrity: present, 1; absent, 0. Inflammatory cell accumulation and fibroblastic proliferation and vascularization: absent, 0; poor, 1; moderate, 2; severe, 3. Additionally, tissue samples were stained with anti-VEGF antibody for vascular endothelial growth factor, and anti-FGF antibody for fibroblast growth factor, and appropriate secondary antibodies were used. The scores were as follows for all proteins’ expression: absent, 0; poor, 1; moderate, 2; and severe, 3.

2.8. Distribution of DFMSCs in the Palatal Tissues

Quantum dot (Qdot)-labeled DFMSCs were analyzed for tissue distribution after the application of DFMSCs with or without hydrogels to determine the homogeneity of cells in palatal wounds. The distribution of Qdot-labeled cells was analyzed using a fluorescence microscope (Nikon Eclipse Ts2, Tokyo, Japan) under 594 nm emission, and DAPI staining was performed to label nuclei.

2.9. Statistical Analysis

Data analysis was performed using GraphPad Prism 8.0 (GraphPad Software). Data are presented as mean (Mean) ± standard deviation (SD) (minimum–maximum) values for each group. Comparisons of data from more than two groups were performed using the ANOVA test (one-way analysis of variance). The unpaired Student t-test was used for comparisons between two groups. Pathological scores were analyzed using the Mann–Whitney U test for two independent samples or the Kruskal–Wallis test for multiple independent samples of nonparametric tests. The results are recorded as Mean ± SD, and p < 0.05 was considered statistically significant.

3. Results

3.1. Dental Follicle Mesenchymal Stem Cells Showed Mesenchymal Stem Cell Characteristics

The third-passage cells exhibited a fibroblast-like morphology in the plastic flasks. The fluorescence intensity of the positive markers of the third-passage cells was over 95% (CD29: 97.6%; CD105: 96.1%; CD73: 99.3%; CD90: 98.7%), while that for the negative markers was lacking (CD14: 1.3%; CD34 0.8%; CD45: 1.7%; HLA-DR: 0.7%). DFMSCs showed potential to differentiate into three lineages (Supplementary Figure S3).

3.2. Quantum Dot-Labeled Cells

DFMSCs were labeled with a quantum dot (Qdot) fluorescent dye for intracellular tracking or to visualize the cell distribution in the hydrogels. The cells showed over 60% positive signal for Qdot in fluorescence imaging (Supplementary Figure S4).

3.3. PRP Quality Tests

In this study, we developed a lyophilization process to preserve the viability of DFMSCs before adding them to the hydrogel. PRP was isolated from the rats’ cardiac blood samples with the lyophilization solution of DFMSCs before producing the hydrogel containing or not containing MSCs for use in an experimental palatal defect animal model. Samples with platelet counts between 1.0 and 1.5 × 10^3^/microliter and >1500 pg/mL PDGF and VEGF in the growth factor analysis were used to prepare MSCs to be incorporated into the biomaterial. The platelet count was within 1.15 × 10^3^ ± 0.12 × 10^3^, VEGF was within 1.948 ± 416 pg/mL, and PDGF was within 3.624 ± 810 pg/mL.

3.4. Characterization of the Hydrogels

The FTIR spectra of the DFMSC-loaded and unloaded hydrogels are presented in Figure 1B. The characteristic absorption bands of the catechol–chitosan backbone were preserved in both samples, indicating that the overall chemical structure of the hydrogel matrix remained unchanged after cell loading. However, owing to the protein- and lipid-rich composition of DFMSCs, an increase in absorption intensity was observed in the Amide I (~1650 cm^−1^) and Amide II (~1540 cm^−1^) regions in the cell-loaded hydrogels. In addition, an increased intensity in the C–O–C stretching vibration region (~1100–1000 cm^−1^) was detected, which can be attributed to the carbohydrate components of the cells. Specifically, the enhanced Amide I and Amide II bands originate from proteinaceous cellular constituents, while the increased C–O–C stretching intensity reflects carbohydrate-containing structures associated with DFMSCs. These changes represent variations in peak intensity rather than the formation of new functional groups within the hydrogel matrix. Overall, comparison of the FTIR spectra of loaded and unloaded hydrogels revealed intensity enhancements associated with cellular components, supporting the successful incorporation of DFMSCs into the hydrogel structure.

SEM images of the unloaded hydrogel revealed a homogeneous, film-like surface with no distinct particulate or crystalline structure. After DFMSCs were loaded, numerous particles were observed as bright spots on the surface, the surface became rough, and a tendency for particles to cluster in some areas was noted. In other words, while drug loading was successful, the surface distribution exhibited a heterogeneous appearance at the microscale (Figure 2A,B).

To quantify these observations, image processing-based particle counting was performed. This analysis yielded an average particle count of 18.5, a standard deviation of 12.4, and a coefficient of variation of 67%, with a minimum of 0 and a maximum of 58. These findings indicated that the loading was heterogeneous at the microscopic scale. Figure 2C shows the grid-based homogeneity heatmap obtained from the particle counting results.

Since DFMSCs were labeled with the quantum dot fluorescent marker before being added to the hydrogels, fluorescence was obtained at a 594 nm wavelength through confocal microscope imaging. After hydrogel production with DFMSCs was completed, lyophilized hydrogels were incubated with DMEM for 2 h to expose the cells, and the samples were frozen and sectioned. Confocal microscope analysis showed that the cells in the hydrogels were homogeneously distributed (Figure 2D).

The swelling rate of the unloaded hydrogels, which started at 1538% in the first 10 min, reached 2160% at 180 min and 2282% at 720 min. No substantial change was observed between 12 and 24 h, indicating that the equilibrium swelling value was achieved at approximately 12 h. DFMSC-loaded hydrogels exhibited lower swelling capacity. The swelling rate of DFMSC-loaded hydrogels, which was 875% at 10 min, reached 1223% at 180 min and 1625% at 720 min. With the swelling value being 1624% at 24 h, it was understood that DFMSC-loaded hydrogels reached the equilibrium swelling value in 6 h. These results indicate that DFMSCs partially fill the internal free volume of the polymer network, limiting the water uptake capacity and reducing the swelling behavior (Figure 3A and Table 1).

Degradation tests revealed that the presence of cells significantly affected the structural stability of the hydrogel. While almost no degradation was observed in unloaded hydrogels (0.00–0.01%) during the test period, a gradual degradation trend was observed in DFMSC-loaded hydrogels. Degradation began at 4.7% in DFMSC-loaded hydrogels at 10 min and reached 21.9% at 24 h. This behavior indicates that cell-associated biological activity, rather than the mere physical presence of cells, contributes to hydrogel degradation. Specifically, biomolecules secreted by DFMSCs, including enzymes and metabolites, together with localized microenvironmental changes within the hydrogel matrix, facilitate polymer network relaxation and degradation, reflecting a controlled degradation mechanism sensitive to biologically mediated stimuli (Figure 3B).

Mucoadhesion tests revealed a clear enhancement in adhesion performance following catechol modification. Although the chitosan hydrogels are initially adherent, the number of attached samples declined rapidly over time: seven remained at 30 min, four at 60 min, and only one at 90 min. After 230 min, none of the hydrogels remained adhered, confirming the limited mucoadhesive capacity of native chitosan. In contrast, catechol-modified hydrogels showed markedly enhanced mucoadhesion. All samples (8/8) remained adhered up to 200 min, after which only one hydrogel detached; the remaining seven stayed attached until the end of the experiment, corresponding to an 88% overall mucoadhesion performance (Figure 3C). This improvement reflects strong noncovalent interactions between catechol groups and mucosal glycoproteins, demonstrating the substantial contribution of catechol modification to mucoadhesive behavior.

The in vitro cumulative release profiles of the hydrogels are presented in Figure 3D. Based on the release kinetics, the data were evaluated in three phases: early (0–20 min), intermediate (20–360 min), and late release (360–1080 min). In the early phase, 1 × DFMSC-loaded hydrogels exhibited minimal release (7.6% at 5–10 min), indicating that most cells were strongly entrapped within the polymer matrix. In contrast, 2 × DFMSC-loaded hydrogels showed a pronounced burst release (42.8% at 5 min; 61.6% at 10 min), reflecting a substantially higher initial release. During the intermediate phase, 1 × DFMSC hydrogels rapidly increased to 61.6% at 20 min and reached 99.9% at 360 min. Although 2 × DFMSC hydrogels exhibited an early burst, their subsequent release became more moderate (64.8% at 20 min; 83.5% at 150 min), ultimately reaching 99.9% at 360 min. In the late phase, both formulations achieved complete release (100%) at 720 min. Because both curves plateaued at 360 min, full DFMSC release was effectively completed within 6 h for both hydrogel groups. The major difference between the release profiles of the two loading ratios was observed in the early phase. This difference can be attributed to two mechanisms: (i) the increased cell concentration enhances diffusion-driven transport, and (ii) at high loading densities, excess cells are unable to fully penetrate into the hydrogel network during incorporation, resulting in a larger fraction remaining near or on the hydrogel surface. These surface-associated cells are therefore released rapidly, giving rise to the observed burst release.

3.5. In Vitro Cytotoxicity Tests

The hydrogels produced in this study were analyzed for cell viability and cytotoxicity. Cell viability tests are crucial for determining the overall health and survival of cells in response to various treatments. A decrease in cell viability of biomaterials indicates a toxic effect, while an increase may indicate a protective or stimulatory effect. In the present study, when live cells were stained with CalceinAM (green) and apoptotic cells with Ethidium Homodimer-1 “EthD-1” (red), over 90% live cells and less than 10% dead cells were observed. These results indicate that the hydrogels had no toxic effects (Figure 4A,B). Consistent with these results, the MTT assay results showed similar viable cell ratios in control cells (93.2 ± 4.1%) and cells cultured with the hydrogels (89.6 ± 3.4%) (Figure 4C).

3.6. Viability of DFMSCs After Dehydration of Hydrogels

The results showed that 7 days after production, 81.8 ± 5.6% of the cells were viable after dehydration of the hydrogels, compared to 93.4 ± 3.8% of the non-lyophilized DFMSCs (Figure 4D–F). In addition, the percentage of viable cells was 72.3 ± 3.4% after dehydration of the hydrogels 5 months after production (Figure 4G,H).

3.7. Lyophilized Hydrogels Loaded with DFMSCs Rapidly Healed Palatal Wounds

Palatal wound areas were measured with a ruler to calculate the wound contraction on days 7 and 14 in rats with a palatal wound model. The percentage of wound contraction was determined by subtracting the wound area from the first day’s wound area. Group 3 (57.6 ± 7.1%), Group 4 (67.5 ± 6.2%), Group 5 (72.0 ± 7.8%), and Group 6 (80.3 ± 7.9%) showed significant wound contraction compared to Group 1 (49.3 ± 5.2%) on day 7 (p < 0.05). In addition, the wound closure percentage was significantly higher in Group 6 compared to Groups 4 and 5 on day 7 (p < 0.05). Group 4 (81.5 ± 5.7%), Group 5 (88.3 ± 6.1%), and Group 6 (94.3 ± 7.0%) showed significant high wound contraction compared to Group 1 (69.8 ± 4.7%), Group 2 (69.0 ± 4.2%), and Group 3 (73.0 ± 3.9%) on day 14 (p < 0.05). The wound closure percentage was significantly higher in Group 6 compared to Group 4 on day 14 (p < 0.05). The results showed that the wound closure rate increased with the DFMSC dose in the hydrogel (Figure 5).

3.8. Histopathology

The epithelial integrity in the palatal wounds in the groups in which hydrogels loaded with DF-MSCs were applied (Group 5 and Group 6), in the group in which hydrogel loaded with lyophilization solution alone was applied (Group 3), and in the group in which lyophilized DF-MSCs were applied (Group 4) was significantly higher than that in the palatal wound group (Group 1) and the group with palatal wounds and hydrogel (Group 2) (p < 0.05). Additionally, ulceration was significantly decreased in the groups in which hydrogels loaded with DF-MSCs were applied (Groups 5 and 6), in the group in which hydrogel loaded with lyophilization solution alone was applied (Group 3), and in the group in which lyophilized DFMSCs were applied (Group 4) compared with that in the palatal wound group (Group 1) and the group with palatal wounds and hydrogel (Group 2) (p < 0.05). The fibroblastic proliferation was severe in Groups 1 and 2, moderate in Groups 3 and 4, and weak in Groups 5 and 6. The statistical analysis showed that fibroblastic proliferation significantly decreased in Group 3 (1.66 ± 0.52) and Group 4 (1.51 ± 0.52) compared to Group 1 (2.67 ± 0.51) and Group 2 (2.50 ± 0.54) (p < 0.05). The fibroblastic proliferation was significantly decreased in Group 5 (0.51 ± 0.54) and Group 6 (0.33 ± 0.52) compared to Groups 1, 2, 3, and 4 (p < 0.05). Ulcers were significantly decreased in Group 3 (0.17 ± 0.40), Group 4 (0.00 ± 0.00), Group 5 (0.00 ± 0.00), and Group 6 (0.00 ± 0.00) when compared with Group 1 (1.00 ± 0.00) and Group 2 (0.83 ± 0.41) (p < 0.05). Inflammatory cell accumulation was significantly decreased in Group 3 (1.83 ± 0.41), Group 4 (1.50 ± 0.55), Group 5 (0.67 ± 0.52), and Group 6 (0.17 ± 0.41) compared to Group 1 (2.67 ± 0.52) and Group 2 (2.50 ± 0.54) (p < 0.05). Vascularization was significantly decreased in Group 3 (1.50 ± 0.55), Group 4 (1.33 ± 0.52), Group 5 (0.50 ± 0.55), and Group 6 (0.17 ± 0.41) compared to Group 1 (2.50 ± 0.55) and Group 2 (2.33 ± 0.52) (p < 0.05). The healing scores of wounds were significantly higher in Group 5 (2.33 ± 0.52) and Group 6 (2.84 ± 0.41) than in Group 1 (0.33 ± 0.52), Group 2 (1.00 ± 0.63), Group 3 (1.50 ± 0.55), and Group 4 (1.67 ± 0.82) (p < 0.05). Although there was no statistically significant difference in wound healing scores between Groups 5 and 6, Group 6 had higher levels of healing parameters than Group 5 and the other groups (Figure 6 and Figure 7).

FGF protein expression was significantly higher in Group 1 (2.33 ± 0.82), Group 2 (2.50 ± 0.55), and Group 3 (2.83 ± 0.41) than in Group 4 (2.16 ± 0.75), Group 5 (0.67 ± 0.82), and Group 6 (0.33 ± 0.52) (p < 0.05). VEGF expression was significantly higher in Group 1 (2.83 ± 0.41), Group 2 (2.67 ± 0.52), Group 3 (2.50 ± 0.55), and Group 4 (2.33 ± 0.52) than in Group 5 (0.33 ± 0.52) and Group 6 (0.17 ± 0.41) (p < 0.05) (Figure 7).

3.9. Distribution of Qdot-Labeled DFMSCs in Palatal Wounds

To visualize the distribution and persistence of DFMSCs in palatal wounds after the application of cells with or without hydrogels, we performed fluorescence microscopy analysis on days 3 and 14. The first three days are important for visualizing the distribution of cells, while day 14 is important for determining the persistence of cells in the wound site [39]. On day 3, while DFMSCs alone showed accumulation in certain areas in the wound site, DFMSCs transferred with the hydrogel showed a homogeneous distribution. At day 14, DFMSCs persisted at the wound site at approximately 25–30% when transferred with hydrogel, but DFMSCs transferred alone persisted at less than 10% in the wound site. The results demonstrated that DFMSCs with hydrogels could be homogenously distributed in the wound site and persisted for a long time (Figure 8).

4. Discussion

Palatal wounds resulting from trauma, tumors, gingival grafts, and other procedures negatively impact quality of life. Palatal wounds impair functions such as mastication and deglutition, while inflammatory responses at the wound site pose life-threatening risks in severe cases. Among surgical procedures, the use of a free gingival graft and a connective tissue graft from the palate are the most common applications, but palatal harvesting causes significant discomfort in the palatal region [40,41]. Acellular dermal matrix, platelet-rich derivatives, amniotic membrane, and hydrogels are some of the therapeutic agents used in the treatment of palatal wounds [41]. In this study, we aimed to develop a ready-to-use therapeutic product that would accelerate wound healing and also achieve scarless healing and full regeneration for application in palatal wounds.

In the present study, we developed a lyophilized hydrogel loaded with MSCs for clinical application in dentistry. Although hydrogels in gel form or lyophilized form and MSCs in fresh form can be used separately in clinical applications, the main problem in practice is the freezing and thawing of cells. Therefore, in this study, DFMSCs were loaded before lyophilization of the hydrogel, and finally, the entire product was lyophilized and produced ready for clinical use. MSCs derived from dental tissues can be easily isolated, have a rapid doubling time, and have a high regenerative potential compared to other MSCs, such as bone marrow MSCs or adipose tissue MSCs. The most potent dental MSCs are DFMSCs, which can differentiate into multiple lineages: epithelial, muscle, bone, cartilage, and neuronal cell types [42]. In this study, we analyzed the viability of freshly prepared DFMSCs and hydrogel-lyophilized DFMSCs. The results indicate that the viability of hydrogel-lyophilized DFMSCs after rehydration is similar to that of freshly prepared DFMSCs, indicating that cell viability can be maintained after hydrogel lyophilization. Additionally, SEM analysis showed that DFMSCs, which were targeted to be released during hydrogel degradation, produced a homogeneous distribution within the hydrogel. One of the main advantages of lyophilized MSCs over cryopreserved cells is their long storage stability. This is crucial not only for preserving the MSCs’ paracrine factors but also for ensuring that the cells maintain their viability and differentiation ability after transplantation into the injured tissue. Therefore, in this study, DFMSCs were blended with a special preservative solution containing trehalose, PRP, and DMSO, which maximized their viability before being added to the hydrogels.

From a mechanistic perspective, DFMSCs may have been protected from osmotic stress and ice crystal formation during lyophilization owing to the lyophilization solution (PRP + DMSO + Trehalose) used. PRP contains a high percentage of albumin, approximately 60%. There is evidence that albumin protects cells against crystal formation and osmotic stress during lyophilization. A study reported that human serum albumin (HSA) and bovine serum albumin (BSA) are commonly used adjuvants in the freeze-drying of cells, and that albumin from both sources enables the storage of biomaterial samples or cells at room temperature in cryopreservation media [43]. Similarly, trehalose is a commonly used lyopreservative in freeze-drying processes [23]. DMSO, on the other hand, is a good crystalline inhibitor used in the cryopreservation of cells. It is highly probable that the PRP + DMSO + trehalose combination we used protected the DFMSCs during both the freezing process (pre-treatment) and the lyophilization process (drying).

Because the oral mucosa is a challenging site for hydrogel fixation, an important outcome of this study is the demonstration that catechol modification markedly enhances the mucoadhesive properties of the hydrogels. Mucoadhesion tests showed that, while unmodified chitosan hydrogels rapidly detached from the mucosal surface, catechol–chitosan hydrogels exhibited substantially prolonged adhesion. These findings confirm that catechol functionalization significantly strengthens hydrogel–mucosa interactions, enabling more stable localization within the oral environment. Another critical parameter evaluated in this study was the release profile of the embedded cells. Notably, hydrogels loaded with an optimal number of DFMSCs exhibited a prolonged and controlled release extending up to 6 h, in contrast to the immediate dispersion expected with direct cell application. This sustained release behavior suggests that delivering cells within a hydrogel matrix, rather than administering them alone, offers significant advantages by ensuring gradual cell availability and improved retention at the target site.

Beyond enhanced macroscopic adhesion, the wet-adhesion chemistry of catechol–chitosan functional groups further explains the improved performance of the hydrogel within the palatal environment. Chitosan is widely used in buccal drug delivery systems due to its biocompatibility, biodegradability, and intrinsic antibacterial activity, all of which are advantageous for palatal wound healing. Its mucoadhesive nature allows prolonged residence at the application site. However, native chitosan exhibits limited adhesion under physiological conditions. Catechol modification significantly enhances the mucoadhesive performance of chitosan. Catechol moieties are capable of forming multiple interaction types under aqueous conditions, including hydrogen bonding with mucin glycoproteins, coordination interactions with metal ions naturally present in tissues, and oxidation-mediated covalent bonding with nucleophilic groups such as amines and thiols on the tissue surface. These catechol-mediated interactions are particularly effective in moist environments, where conventional mucoadhesive polymers often fail to maintain stable adhesion. In the context of palatal wound healing, such interactions enhance hydrogel retention on wet mucosal tissues, thereby maintaining close contact between the biomaterial, the wound bed, and the delivered DFMSCs. This prolonged residence time supports localized biological activity, effective cell retention, and modulation of the early wound-healing microenvironment [20,30,34]. A schematic illustration of the catechol-mediated wet tissue adhesion mechanism is provided in Supplementary Figure S5.

The negligible degradation observed in unloaded catechol–chitosan hydrogels over 24 h indicates a high short-term structural stability of the genipin-crosslinked polymer network under physiological oral conditions (pH 6.8, 37 °C). In the absence of cells, degradation is governed primarily by passive hydrolytic processes, which are limited within a short time. All degradation experiments were performed in the same buffered medium, ensuring identical external ionic strength for unloaded and DFMSC-loaded hydrogels. Accordingly, the increased degradation observed in DFMSC-loaded hydrogels cannot be attributed to differences in the bulk ionic strength of the surrounding medium. Instead, this behavior is likely driven by biologically mediated mechanisms, including cell-secreted enzymes, metabolites, and localized microenvironmental changes induced by DFMSCs and the cell-preserving formulation, which collectively facilitate polymer network relaxation and degradation.

Palatal wound healing was assessed by analyzing tissue sections for inflammatory cell accumulation, fibroblastic proliferation, vascularization, ulceration, and healing scores. The results of this study demonstrated significantly higher regeneration in palatal wounds in the DF-MSC-loaded hydrogel-treated groups. One of the fundamental stages of wound healing is the inflammation phase, which is a transition period between wound hemostasis and proliferation from the moment of tissue damage to the recovery of the barrier and other functions of the tissue. However, if acute inflammation is insufficient and prolonged, the inflammation becomes pathogenic, leading to impaired interaction between different cell types, such as fibroblasts and other tissue cells. In this case, inflammation also causes fibroblastic proliferation, resulting in delayed wound healing and scarring [44]. It is well known that MSCs achieve wound healing without scarring [45]. During wound healing, MSCs secrete proangiogenic growth factors, facilitating rapid wound healing. Vascularization increases in the early stages of wound healing with the effect of growth factors, while growth factors decrease as healing is completed [46]. FGF is a growth factor that promotes fibroblastic proliferation and also promotes wound tissue formation, resulting in scar healing [47]. In this study, decreased inflammatory cell accumulation and ulceration were observed in the palatal tissues of groups treated with DF-MSC-loaded hydrogels. In addition, FGF and VEGF expressions were significantly decreased in the palatal tissues of groups treated with DFMSC-loaded hydrogels. The data demonstrated that hydrogels loaded with DFMSCs accelerated healing in palatal wounds. Although no significant difference was observed between the two different DFMSC doses, high-dose DFMSC loading into the hydrogels dose-dependently enhanced healing. These results suggest that loading hydrogels with DFMSCs during fabrication and then lyophilizing them better enhances palatal wound healing.

There are numerous studies on the use of various biomaterials, such as bioceramics, alginate, chitosan, hydrogels, cell sheets, nanoparticles, and 3D printing, in tissue engineering and inflammation. MSCs treated with biomaterials can survive better than MSCs transferred alone. MSC transplantation is a safe and effective cell regeneration therapy, but the reparative effect of direct cell transplantation is unstable [48]. Vector materials have their own advantages and disadvantages when combined with MSCs, but currently, MSCs are combined with biomaterials just before application in vivo [49,50]. Bolinas et al. reported that injectable hydrogels loaded with MSCs are stable for 14 days [51]. In this study, we analyzed the viability of cells in DFMSC-loaded hydrogels 5 months after production and showed that the viable cell percentage was approximately 71%.

The incorporation of MSCs into biomaterials is a current and important topic in clinical medicine, particularly in dentistry and orthopedics. The MSCs loaded with biomaterials are known to significantly impact tissue repair, not only through the biomaterial’s own effect on tissue repair but also through the versatile differentiation potential of their cellular content and their strong paracrine capacity, which is a crucial mechanism supporting tissue repair. Stimuli from the extracellular environment influence the proliferation and differentiation properties of MSCs. Therefore, accelerating the repair of defective tissues creates a synergistic effect through the combination of biomimetic materials, the potential of MSCs to differentiate into tissue cells, and the influence of paracrine factors secreted by MSCs [52]. Recently, researchers have begun to investigate the synergistic effects of combined MSC–biomaterial therapies, where the biomaterial acts as a scaffold to protect MSCs and provides physiologically relevant physicochemical signals that can direct MSCs’ immunomodulatory behavior. A number of factors limit the clinical application of MSC therapies, including availability, the uncertainty regarding the cell numbers per dose, the low survival rate, biocompatibility, efficacy, and cost. Preclinical studies to date suggest that biomaterials may help overcome these limitations [48].

From a periodontal clinical perspective, the accelerated and more physiologically balanced healing response observed with DFMSC-loaded lyophilized catechol–chitosan hydrogels holds considerable translational relevance. Autogenous palatal graft harvesting remains the gold standard for soft-tissue augmentation procedures such as connective tissue grafting and free gingival grafting. However, it is frequently associated with postoperative pain, bleeding, delayed epithelialization, and donor site ulceration, all of which negatively affect patient comfort, treatment acceptance, and overall morbidity. The marked reductions in inflammation, fibroblastic hyperproliferation, and ulceration demonstrated in the present study indicate that this bioactive hydrogel system may alleviate these well-documented complications. The improvements in epithelial integrity and wound contraction further suggest that MSC-loaded hydrogels may act as a biologically active dressing capable of modulating early wound-healing dynamics toward regenerative rather than reparative outcomes. Such an approach may shorten recovery time following mucogingival surgery, enhance postoperative comfort, and ultimately contribute to improved donor site management in periodontal plastic surgery. The ready-to-use, lyophilized format of this construct also provides a clinically feasible alternative that overcomes the logistical limitations associated with fresh MSC delivery, thereby increasing the likelihood of successful clinical translation.

Nevertheless, several limitations of the present study should be acknowledged. Although the DFMSC-loaded lyophilized hydrogels demonstrated favorable immunomodulatory and regenerative effects in palatal wound healing, the findings were derived from an experimental model with a limited observation period and may not fully reflect the complexity of long-term host immune interactions in clinical settings. In addition, the use of a single MSC source and hydrogel formulation restricts direct comparison with alternative cell types or biomaterial strategies. Therefore, further studies incorporating extended follow-up, comprehensive immunological profiling, and controlled clinical trials are warranted to substantiate the translational applicability of this approach. In addition, although the catechol functionalization of chitosan was qualitatively confirmed by UV-Vis spectroscopy through the characteristic absorbance peak at 280 nm, quantitative determination of the catechol conjugation degree using NMR or calibrated UV-Vis analysis was not performed in the present study. Furthermore, thermal characterization, such as thermogravimetric analysis, was not included because the primary focus of this work was biological performance and regenerative outcomes under physiological conditions. Future studies will incorporate quantitative polymer characterization and comprehensive thermal analyses to further elucidate the physicochemical properties of the hydrogels and to better correlate material chemistry with biological efficacy.

5. Conclusions

In conclusion, the present study demonstrates that hydrogels loaded with mesenchymal stem cells during fabrication represent a promising therapeutic strategy for accelerating palatal wound healing. The lyophilized DFMSC-loaded hydrogel system preserved high cell viability following rehydration, with more than 80% viable cells observed at 7 days after production and over 70% viability maintained even after 5 months of storage. By combining controlled cell delivery, enhanced mucoadhesion, and immunomodulatory effects, this approach promoted a more balanced healing response characterized by reduced inflammation and improved tissue regeneration. Collectively, these findings highlight the potential of ready-to-use, lyophilized DFMSC-loaded hydrogels as a clinically feasible and translationally relevant platform for improving palatal donor site healing in periodontal applications.

6. Patents

The lyophilized hydrogel containing dental follicle-derived mesenchymal stem cells investigated in this study was developed by the authors. A patent application related to this biomaterial may be filed, and the authors retain the intellectual property rights associated with its development.