Development of Antimicrobial Wound Healing Hydrogels Based on the Microbial Polysaccharide Pullulan

Abstract

Microbial polysaccharides are promising components for wound-care products. This study reports the development of wound-healing antimicrobial hydrogels, based on pullulan from Aureobasidium pullulans, combined with mesenchymal cell-derived conditioned medium. Structural characterization of pullulan was confirmed by FTIR and NMR. Twenty-three formulations containing pullulan, chitosan, gelatin, citric acid, and antimicrobial agents were prepared. Physicochemical screening identified optimal hydrogels: No. 22 (1.2% pullulan, 1.2% chitosan, 0.2% citric acid, 2.4% gelatin, 0.1% conditioned medium, 0.4% glutaraldehyde) and No. 23 (2.4% pullulan, no chitosan, the remaining components identical to those in No. 22). Both exhibited pH values of 5.34 and 5.49, moisture content of 92%, swelling capacities of 175% and 213%, and dynamic viscosity between 58–120 mPa·s. Cytotoxicity testing with human mesenchymal stem cells showed no significant toxicity, with both hydrogels supporting cell adhesion and proliferation. Antimicrobial assays demonstrated inhibitory activity against Staphylococcus aureus and Escherichia coli for both formulations; only hydrogel No. 23 inhibited Pseudomonas aeruginosa. In vitro scratch assays revealed that hydrogel No. 23 significantly promoted fibroblast migration, achieving 30.25% scratch closure after 24 h. The developed formulations combine favorable physicochemical properties with antimicrobial efficacy and regenerative potential, supporting further evaluation as advanced wound-healing and anti-burn dressings.

1. Introduction

In recent years, increasing attention has been paid to the development of biocompatible and biodegradable wound healing and antimicrobial agents based on natural biologically active substances, including microbial polymers [1,2]. Among the many newly developed wound dressings aimed at optimizing the regenerative process and improving patients’ quality of life, hydrogels appear to be particularly promising. They are often based on microbial polysaccharides, which possess unique physical and chemical properties that can be precisely adjusted depending on the intended use. Hydrogels are three-dimensional polymer networks that can hold a significant amount of water or biological fluids due to their structural porosity and the presence of hydrophilic functional groups [3]. Compared to traditional oil- and fat-based ointments or even other polymeric dressings, hydrogel dressings offer several advantages. These include moisture permeability, high sorption capacity, removal of excess exudate, barrier function against secondary infection, mechanical stability, adaptability to wounds with irregular surfaces, prevention of desiccation, wound surface cooling, and pain relief [4,5].

Wound healing is a complex biological process involving tissue growth and regeneration. It progresses through a series of interrelated stages where various cellular and matrix components work together to restore the integrity of damaged tissue and replace lost tissue [6].

The benefits of a moist environment in wound healing were first noted by Winter and have since been supported by numerous studies [7].

Natural materials used for the production of medical hydrogels include collagen, elastin, gelatin, chitosan, dextran, pullulan, hyaluronic acid, alginate, silk fibroin, and glycosaminoglycans. A significant advantage of biopolymer-based hydrogels is the availability and low cost of raw materials.

Microbial polysaccharides can serve as a safe and economical matrix in the development of low-allergenicity hydrogels. These components can retain moisture, stimulate cell adhesion, and accelerate wound healing [8].

According to the National Institute of Health, the use of microbial polysaccharides in pharmaceutical products has contributed to a significant reduction in adverse effects associated with the use of synthetic components. The global market for microbial polysaccharides in the pharmaceutical sector is estimated to reach $2.7 billion by 2026, reflecting their growing relevance in medical innovations [9].

Pullulan, a microbial polysaccharide widely used in biomedicine, is a promising matrix for wound-healing gels due to its antioxidant activity, acceleration of healing processes, non-immunogenicity and environmental safety. However, its mechanical strength is limited, necessitating its combination with other polymers to improve stability and durability of the polymer mesh, thus creating more favorable conditions for epithelialization [10].

Chitosan, a cationic polysaccharide capable of forming numerous ionic and hydrogen bonds, can consolidate the polymer network and increase its mechanical strength [11]. When combined with gelatin, it can form a double network structure that increases the viscoelasticity of gels and their turgor—properties that are desirable in regenerative medicine [12].

Although synthetic polymers offer greater resistance to sterilization, degradation and the attack of microorganisms, the intrinsic activity and synergistic effects of natural polymers make them preferable in medical applications. Their functional groups enable chemical modifications, allowing fine-tuning of biological, rheological, and physicochemical properties of the resulting hydrogels [13].

Recent advances also highlight the large therapeutic potential of mesenchymal cell-derived conditioned media [14]. These media accelerate tissue regeneration and promote cell migration and proliferation through a broad spectrum of growth factors, exosomes, cytokines, and other bioactive molecules. The polymer matrix can support the controlled release of active substances, while the conditioned media provide signaling molecules to enhance therapeutic efficacy [15,16].

The aim of this work is to develop a wound-healing hydrogel based on the microbial polysaccharide pullulan. The main objectives of the study are: (a) characterization of the physicochemical properties of pullulan; (b) development and characterization of hydrogel formulations; and (c) analysis of the wound-healing properties of the hydrogels in vitro.

An innovative antimicrobial hydrogel composition for topical application with improved therapeutic potential has been developed. The combination of microbial pullulan and cell-derived conditioned medium ensured high moisture retention, biocompatibility and favorable rheological properties. The efficacy of the experimental hydrogel formulations was demonstrated in vitro using fibroblast models.

2. Materials and Methods

2.1. Polysaccharide Extraction

In this study, pullulan polysaccharide was used, produced by the yeast-like fungus Aureobasidium pullulans C7 (GenBank accession number: OR864236).

A. pullulans C7 was cultivated in Czapek-Dox broth (Merck, Germany) on an orbital shaker-incubator ES-20 (Biosan SIA, Riga, Latvia) at 150 rpm for 5 days at 25 °C. The culture broth was separated from the biomass by centrifugation using an RS-6MC Dastan centrifuge (Kyrgyzstan) at 6000× g for 20 min.

The polysaccharide was precipitated from the supernatant by mixing with cold (5 °C) 96% ethanol in a 2:1 (v/v) ratio and dried at room temperature for 2–3 days. The yield of crude pullulan was 19.05 ± 0.48 g/L.

2.2. IR Spectroscopy

Fourier transform infrared (IR) spectroscopy was performed on a PerkinElmer Spectrum IR spectrophotometer (Waltham, MA, USA) in the range of 450–4000 cm⁻¹. The pullulan sample was first ground to a fine powder, and 1 g of purified powder was transferred to a metal platform for uniform contact with an attenuated total reflectance crystal. Each sample was scanned three times to ensure a high signal-to-noise ratio.

2.3. NMR Analysis

Nuclear magnetic resonance (NMR) analysis was performed at 80 MHz frequency using a benchtop NMR spectrometer (NMR Spinsolve, Magritek GmbH, Aachen, Germany), equipped with an autosampler and Spinsolve software (version 2.2.3). The pullulan sample was dissolved in deuterium oxide (D₂O) at a concentration of 25 mg/mL to suppress the proton signal of the solvent. A total of 0.5 mL of the solution was transferred into a standard 5 mm NMR glass tube. The NMR spectra were recorded at room temperature with the following parameters: accumulation time (AT) = 3.2 s, repetition time (RT) = 30 s, number of scans (NS) = 64, and pulse angle (PA) = 90°.

2.4. Selection of Hydrogel Components

The hydrogel compositions were prepared using the following components: pullulan (isolated from A. pullulans C7), chitosan (ALDRICH, Iceland, molecular weight 190–375 kDa), citric acid (Ph. Eur., 99.5 to 100.5%, AppliChem, Germany), gelatin (gelatina medicinalis, Ph.Eur., 99.5 to 100.5%, AppliChem, Germany, molecular weight 50–100 kDa), glutaraldehyde (25% solution, Sigma-Aldrich, St. Louis, Missouri, USA), purified water (aqua purificata, water purified, Chistayavoda, Kazakhstan, GOST 6709-72), sterile Dulbecco’s Modified Eagle Medium (DMEM, Gibco, USA), selected after cultivation of mesenchymal stem cells (passage number—3, time of culture—5 days), provided by the Medical Centre for Cell Therapy, SEPINEO™ DERM (SEPPIC, France), 1% lidocaine hypochlorite (SANTO, Kazakhstan), 0.05% chlorhexidine bigluconate (Shansharov-Farm, Kazakhstan), chloramine T (supplier AMK-Chemico, manufacturer China) (

Table 1. Components for forming hydrogel.

Ingredient Names	Purpose of Ingredients
pullulan	polymer matrix
chitosan	polymer matrix
SEPINEO™ D.E.R.M.	emulsifier
citric acid	preservative, pH regulator
gelatin	emulsifier
HPLC water	solvent
DMEM	solvent
glutaraldehyde	cross-linker, disinfectant
chlorhexidine	antiseptic
chloramine T	antiseptic

and

Table 2. Composition of the developed hydrogel samples.

Hydrogel Samples	Pullulan (g)	Chitosan (g)	SEPINEO™ D. E. R. M (g)	Citric Acid (g)	Gelatin (g)	Glutaraldehyde (mL)	Lidocaine 1% (mL)	Chlorhexidine (mL)	Chloramine T (mL)	HPLC Water (mL)	DMEM (mL)	Total (mL)
1	0.6				3					100		100
2	0.6			0.03	3			0.5		99.5		100
3	0.6			0.035	5			1		99		100
4	0.6			0.03	3			0.5		99.5		100
5	0.6				5			0.5		99.5		100
6	0.6			0.03	5			0.5		99.5		100
7	0.6				3				0.5		99.5	100
8	0.6			0.03	3				0.5		99.5	100
9	3				3			0.5			99.5	100
10	3			0.03	3			0.5			99.5	100
11	3			0.03	3						100	100
12	3				3						100	100
13	3		3	0.03				0.5		99.5		100
14	1.5	1.5		0.6	3		50	0.5		49.5		100
15	1.5	1.5		0.6	3		50	0.5			49.5	100
16	3			0.5	3	0.5				99.5		100
17		3		0.5	3	0.5				99.5		100
18	1.5	1.5		0.4	3	0.5				99.5		100
19	3				3	0.5				99.5		100
20	3			0.7	3	0.5					99.5	100
21		3		0.7	3	0.5					99.5	100
22	1.2	1.2		0.2	2.4	0.4					99.5	100
23	2.4			0.2	2.4	0.4					99.5	100

).

2.5. Hydrogel Preparation

The 23 hydrogel formulations were designed to systematically vary the concentration of pullulan (0.6–3%), chitosan (1.2–3%), gelatin (2.4–5%), and crosslinking agents. As the solvent, either HPLC water or conditioned DMEM was used depending on the sample composition. For samples 1–6, 13–14 and 16–19 HPLC water was employed; for samples 7–12, 15 and 20–23, conditioned DMEM was used. The goal was to optimize mechanical and biological performance parameters. The hydrogel preparation procedure is illustrated in Figure 1. Gelatin was weighed and left to swell in the solvent (either sterile distilled water or DMEM-conditioned medium) for 30 min. Chitosan was dissolved in a mildly acidic solution of citric acid (pH 5–6) and stirred at 100 rpm for 2 h at room temperature using a magnetic stirrer (MSH-300, Biosan Sia, Riga, Latvia). All components were added gradually under continuous stirring and gentle heating to 38 °C. For chemical crosslinking, 0.4–0.5% glutaraldehyde was added dropwise during the final mixing step and allowed to react for 15 min under stirring at 38 °C, followed by neutralization with 0.1 M glycine solution. Stirring continued until the mass thickened and turned into a gel. The resulting gel mass was poured into sterile Petri dishes for cooling to room temperature.

2.6. Hydrophobicity/Hydrophilicity Testing

To determine the hydrophobicity/hydrophilicity of the gel, a mixing and dilution test was used, which is a modification of the drop test for determining the type of emulsion. Two drops of the hydrogel sample were placed on the surface of a Petri dish at a distance of 2 cm from each other. Then, a drop of water was placed next to one drop, and a drop of oil was placed next to the other. If the hydrogel drop disperses quickly in oil but remains agglomerated in water, the sample is hydrophobic, which corresponds to the ‘water-in-oil’ type. Conversely, dispersion in water and cohesion in oil indicated hydrophilicity (oil-in-water system) [17].

2.7. pH Measurement

The hydrogen index was measured at room temperature (25 ± 1 °C) using a glass pH electrode connected to a calibrated pH meter (pH-15-MI, Russia). Undiluted hydrogel (10 mL) was placed in a clean glass beaker, and the electrode was immersed directly into the sample.

2.8. Moisture Content in Hydrogels

Moisture content was determined gravimetrically. For each sample, 2.0 g of hydrogel was weighed into clean porcelain dishes and left at room temperature for 48 h. The samples were then reweighed using an analytical balance. Moisture content was calculated using Equation (1):

Moisture content (%) = (W1 − W2)/W1 × 100

(1)

where W1 is the initial weight of the sample before incubation, and W2 is the weight of the sample after 2 days of drying.

2.9. Swelling Degree

The degree of swelling of the samples was determined gravimetrically. Samples were pre-frozen at −20 °C and freeze-dried using an INOFD-10SU lyophilizer (Qingdao, China). Dried gels were immersed in distilled water at room temperature until equilibrium swelling was reached.

The swelling degree (S) was calculated using Equation (2):

S = (m (w) − m (d))/(m (d)) (g H₂O/g hydrogel)

(2)

where m (w) is the mass of the wet sample and m (d) is the mass of the dried sample.

2.10. Morphological Characterization

The morphology of hydrogels was studied using scanning electron microscopy (SEM) on lyophilized samples. The samples were fixed to stubs with carbon tape. Quanta 200i 3D scanning electron microscope (FEI Company, Hillsboro, OR, USA) was used for observation at an accelerating voltage of 15 kV and a working distance of 10 mm. Imaging was performed at a high vacuum mode of 6 × 10⁻⁶ Pa.

2.11. Dynamic Viscosity

The dynamic viscosity of hydrogels was studied using a rotational viscometer (Brookfield DVE, Ametek, Berwyn, PA, USA) equipped with a ULA spindle. Measurements were taken at shear rates from 30 to 100 rpm at two temperatures (25 °C and 60 °C), using 100 mL of sample.

2.12. Thermogravimetric Analysis

Thermogravimetric analysis (TGA) was performed to study the stability of the hydrogel prototypes. The samples were previously freeze-dried, then 11.1 g of sample 22 and 10.0 g of sample 23 were used to evaluate the chemical characteristics of the decomposition process. TGA measurements were performed using a Labsys Evo differential thermal analyzer (Setaram, Caluire-et-Cuire, France). The samples were heated at a constant rate of 10 °C/min in the presence of air in the temperature range 30–600 °C. The parameters T₅% and T₁₀% (5 and 10% weight loss of the samples), T_max (maximum decomposition temperature) and residual mass at 600 °C were determined. The data were processed using the instrument’s software and then visualized in Origin 9.0 (OriginLab Corporation, Northampton, MA, USA) [18].

2.13. Antimicrobial Activity

The antimicrobial activity of the hydrogel samples was evaluated using the agar well diffusion method on nutrient agar (Merck, Darmstadt, Germany). The following test cultures were used: Staphylococcus aureus ST228, Escherichia coli 603 and Pseudomonas aeruginosa 853. Microbial suspensions of the test culture were prepared in physiological solution at concentrations corresponding to a turbidity standard of 0.5 McFarland. Each suspension was spread evenly over the surface of a nutrient agar plate and allowed to dry under sterile conditions. Wells with a diameter of 1.5 cm were made using a sterile cork borer, and 1 mL of each hydrogel sample was placed into each well. The plates were incubated for 24 h at 36 °C. Zones of inhibition were measured to evaluate antimicrobial activity.

To determine the minimum inhibitory concentration (MIC) of the hydrogel prototypes, the agar dilution method was used according to Clinical and Laboratory Standards Institute (CLSI) guidelines [19]. The method was modified to adapt it for polymer systems. Hydrogels in concentrations from 10 to 80% were added to nutrient agar (Difco Laboratories, Detroit, MI, USA) cooled to 45 °C. The mixtures were thoroughly mixed and placed in sterile Petri dishes. A suspension adjusted to 0.5 McFarland standard and further diluted to 1 × 10⁵ CFU/mL was prepared from fresh bacterial cultures of S. aureus ST228, E. coli 603, and P. aeruginosa 853. 1 mL of the cell suspension was applied onto the surface of agar plates containing different concentrations of hydrogel in the composition and evenly spread over the entire surface. The plates were incubated at 36 °C for 24 h. The control cups were: growth control (nutrient medium without hydrogels), sterility control (nutrient medium without hydrogels and without microorganisms), positive controls (ciprofloxacin for P. aeruginosa, vancomycin for S. aureus, ampicillin for E. coli).

MIC was defined as the lowest hydrogel concentration that completely inhibited visible microbial growth on the agar surface. All experiments were performed in triplicate.

2.14. Effect of Hydrogels on Biofilm Formation

The antibiofilm activity of hydrogel samples No. 22 and No. 23 was evaluated using the method of Saising et al. with modifications for S. aureus ST228 and P. aeruginosa 853. The test samples were added to trypticase soy broth (TSB, 2% glucose; Difco Laboratories, Detroit, Michigan, USA) at concentrations of 30% and 50%. Bacterial cultures at a concentration of 10⁵ CFU/mL and the following variations were seeded into the wells of a 24-well plate (Costar, Glendale, Arizona, USA): S.aureus ST228, P. aeruginosa 853, S. aureus ST228 + P. aeruginosa 853. The total volume of liquid in the well was 3 mL (1 mL of bacterial suspension + 2 mL of TSB with hydrogels). No hydrogels were added to the control wells. Incubation was carried out under static conditions at 36 °C for 24 h [20].

A day later, the supernatant was removed from the wells, and the wells were washed with a phosphate-buffered saline (PBS). The cells adhering to the bottom of the wells were stained with 0.1% crystal violet solution (Sigma-Aldrich, St. Louis, Missouri, USA). Then the dye was removed, the materials were washed with sterile distilled water and dried at 30 °C. The dye bound to the biofilm was extracted with 95% ethanol. Optical density (OD) was measured on a LEKI SS1207 UV spectrophotometer (MEDIORA OY, Helsinki, Finland) at 570 nm. The level of biofilm formation was assessed according to the following criteria: OD < 0.1—negative, 0.1 ≤ OD < 1—slightly positive, OD ≥ 1—positive [20].

2.15. Cytotoxicity

Given that the development of a pharmaceutical product requires comprehensive evaluation of its biocompatibility, the cytotoxicity of the experimental hydrogels was assessed using the direct contact method in accordance with ISO 10993-5 [21]. Human mesenchymal stem cells (MSCs), known for their high adhesion capacity, were used as the cell model [18].

MSCs (4.3 × 10⁵ cells/well) were seeded in 12-well plates on DMEM supplemented with 10% bovine serum and hydrogels No. 22 and No. 23: 10%, 30%, 50%, 70%. The total volume of the suspension in each well was 1 mL. The cells, in the presence of hydrogels, were incubated for 3, 5 and 7 days at 37 °C and 5% CO₂ [22]. Cell viability was assessed every 3, 5, and 7 days of incubation using the Trypan Blue Exclusion Test, according to the Strober protocol [23]. Prior to incubation, cell viability was 94 ± 1%. Cell counting and viability calculations after staining were performed using the Countess II automatic cell counter (Thermo Fisher Scientific, Waltham, MA, USA).

2.16. Scratch Assay (Wound Healing In Vitro)

Wound healing was assessed using the scratch analysis method in vitro on a culture of donor skin fibroblasts provided by the Medical Centre for Cell Therapy (donor information is not disclosed, donor ID: SD). Fibroblasts were cultured in 75 cm² cell culture flasks with slanted necks and ventilated caps (NEST, Wuxi, China), using high-glucose DMEM (4.5 g/L; Gibco, Waltham, MA, USA) supplemented with 10% FBS (Gibco, Waltham, MA, USA). Prior to use, culture media were warmed to 36.6 °C in a water bath (BioSan, Riga, Latvia), and all liquids were dispensed using sterile, individually packaged serological pipettes (NEST, Wuxi, China). Cells were incubated in a Binder CB150 CO₂ incubator (Tuttlingen, Germany).

Cells were retrieved from cryopreservation and thawed in warmed DMEM. After adherence and formation of a confluent monolayer, a linear wound (“scratch”) was introduced by gently scraping the cell monolayer with a sterile plastic loop tip, without damaging the plastic surface. Detached cells were removed by rinsing with sterile physiological saline. Hydrogel samples were then applied. Microscopic fields were marked with a permanent marker to ensure consistent positioning during observation. Cell migration into the scratch area was monitored after 3, 6, and 24 h using an inverted microscope (Olympus CKX53, Evident, Tokyo, Japan). For each test condition, seven replicates were performed. Wound closure was quantified by measuring the scratch area before and after 24 h of incubation.

2.17. Statistical Analysis

All measurements were performed in triplicate unless otherwise stated. Data are presented as the mean ± standard deviation (SD). Statistical analysis was conducted using Statistica software version 10.0 (TIBCO Software Inc., Palo Alto, CA, USA).

3. Results and Discussion

3.1. Physicochemical Properties of Pullulan

Microbial polysaccharides such as xanthan, dextran, pullulan have recently found wide application in pharmaceutical and cosmetic preparations. Therefore, the first stage of the study was to obtain the natural polysaccharide pullulan for further inclusion in wound-healing polymer hydrogels. This choice was made because it fully meets the specific requirements for polymers intended for medical use: biocompatibility, non-toxicity, stability, and ability to withstand the sterilization process.

It is known that pullulan is produced by yeast-like fungi A. pullulans. The structure and composition of the polysaccharide may vary depending on the characteristics of the strain and the nutrient substrates provided [24]. In our previous work, the A. pullulans C7 strain produced exopolysaccharide in amounts ranging from 15.08 ± 0.34 to 34.66 ± 1.73 g/L on various nutrient media, which corresponds well to yields reported by other researchers (15–40 g/L) [24,25,26,27].

To enable the use of the isolated pullulan in wound-healing hydrogel formulations, its structure and physicochemical properties were further examined.

Fourier-transform infrared (FTIR) spectroscopy was employed to identify the functional groups and confirm structural characteristics typical of pullulan. The results are presented as a plot of % transmittance versus wavenumber in the range of 450–4000 cm⁻¹ for qualitative interpretation (Figure 2).

FTIR-spectroscopy revealed peaks characteristic of polysaccharides. The broad absorption band in the 3200–3600 cm⁻¹ region corresponds to O–H stretching vibrations, indicating the presence of hydrogen bonds. The depth of this band reflects the degree of structural order. Narrow peaks observed in the 2850–3000 cm⁻¹ region correspond to C–H stretching vibrations, attributed to the carbon framework of glucose residues. The 1000–1200 cm⁻¹ region shows intense bands associated with C–O–C and C–O bond, whose presence indicates the regularity of the maltotriose linkages. The peaks in the 1050–1150 range are particularly pronounced (a smooth decrease of 0.67%), confirming the presence of α-glycosidic bonds. The region between 800–950 cm⁻¹ is characteristic of the overall polymer structure and supports the presence of a linear, unbranched chain.

These results obtained are consistent with the data of other researchers. For instance, the IR study by Victor R. L. Oliveira reported similar absorption bands: 3430–3440 cm⁻¹ (O–H stretching), 2930 cm⁻¹ (C–H stretching), 1150–1030 cm⁻¹ (C–O–C and C–O stretching), and around 950 cm⁻¹ (indicative of α-1,6-D-glycosidic linkages) [28].

Thus, the IR spectrogram confirmed the polysaccharide nature of the obtained substance and its structural similarity to pullulan, composed of α-1,6-linked maltotriose units.

In order to confirm the structure and composition of pullulan molecules, NMR analysis was performed (Figure 3). In the 5.0–5.5 ppm range, weak anomeric proton signals (relative intensity 0.070–0.076) are observed in the NMR spectrum, confirming the presence of both α-1,4- and α-1,6-glycosidic bonds characteristic of the pullulan structure. Additional peaks indicate the presence of protons attached to carbon atoms bearing hydroxyl groups or CH₂ moieties. Importantly, the spectrum shows the absence of peaks between 0 and 2.0 ppm, indicating the absence of aliphatic impurities and methyl impurities. Similarly, the absence of peaks between 6.0 and 9.0 indicates the absence of aromatic structures.

In order to confirm the structure and composition of pullulan molecules, NMR analysis was performed (Figure 3). The peak between 5.1–5.3 ppm corresponds to protons at the anomeric carbon involved in α-1,6-glycosidic linkages (H1), while signals between 4.9 and 5.1 ppm are indicative of α-1,4-glycosidic bonds. Additional peaks indicate the presence of carbon atoms bound to hydroxyl groups or CH₂ moieties. Importantly, the spectrum shows the absence of peaks between 0 and 2.0 ppm, indicating the absence of aliphatic impurities and methyl impurities. Similarly, the absence of peaks between 6.0 and 9.0 indicates the absence of aromatic structures.

Thus, the characteristic chemical shifts and absence of impurity signals confirm the structural purity and polysaccharide nature of the pullulan sample. These findings are in agreement with the results of a study of commercial pullulan from Sigma Chemical Company and Pfanstiehl Laboratories, which showed that pullulan consists exclusively of maltotriose units linked by α-1,4-glycosidic bonds [29].

3.2. Development of Hydrogel Composition

At the next stage of the study, the structurally confirmed pullulan was incorporated into hydrogel formulations intended for wound healing.

A total of 23 hydrogel samples for external application were prepared. These formulations included compounds acting as a polymer matrix, emulsifiers, preservatives, solvents, disinfectants, and antiseptics (Table 2).

Pullulan was selected as the basis for the polymer matrix due to its unique physical and chemical properties. A number of studies confirm its safety, antitumor and anti-inflammatory properties [30,31]. It is known that when pullulan is used as the sole gelling agent in pharmaceutical and cosmetic formulations, its concentration should not exceed 30% by weight, to achieve optimal gel texture, viscosity and stability [30]. In the hydrogels developed in this study, additional gelling agents—gelatin and chitosan—were used alongside pullulan. This combination allowed us to reduce the pullulan concentration to 3%, providing a balance between effective gel formation, desirable consistency, and ease of application.

Chitosan is a cationic polymer, a deacetylated derivative of chitin, which has anti-inflammatory, antioxidant, antimicrobial, fungicidal, antitumor and regenerative properties [32,33,34,35,36,37]. It is soluble in weakly acidic aqueous media, forming positively charged chains upon protonation of amino groups, which weakens interchain interactions and facilitates gel formation. In our study, chitosan was used at concentrations ranging from 1.2% to 3%. It also improves mechanical strength, biodegradability, and adhesion, and is recognized as non-toxic—making it particularly well-suited for wound dressing applications [38].

Gelatin, a denatured form of collagen, is well known for its biocompatibility, biodegradability, and low immunogenicity. It is approved by the U.S. Food and Drug Administration as a safe compound. Due to its gel-forming capacity and biological activity, gelatin was selected as an auxiliary polymer for hydrogel preparation. It is highly soluble in hot water (up to 80 °C in our case) and exhibits thermoreversibility. The gelatin concentration used in our formulations did not exceed 5% of the total volume, which is known to result in gels with good mechanical stability and moderate swelling capacity—important for maintaining form and enabling controlled release of active substances [39,40].

The hybrid polymer network formed by pullulan, chitosan, and gelatin offers multiple advantages due to the synergistic interaction of their individual properties. Pullulan contributes to structural integrity and controlled substance release; chitosan imparts antimicrobial and bioadhesive properties; gelatin enhances elasticity and flexibility. Such combinations are increasingly described in the literature as promising for biomedical applications, particularly in wound dressings and local drug delivery systems [41,42].

Citric acid is widely used in medicine as a pH regulator. It is known that an acidic aqueous environment (pH below 6) is necessary to dissolve chitosan, promoting the protonation of amino groups present in the chitosan structure, making the polymer positively charged and thereby weakening the binding forces between the chitosan chains [43,44].

The concentration of citric acid we have chosen, up to 0.7%, ensures sufficient protonation of amino groups to dissolve chitosan and promotes the formation of a homogeneous gel structure [45,46].

Glutaraldehyde, a bifunctional compound containing two aldehyde groups, is known for its strong crosslinking ability due to covalent bonding with amino and hydroxyl groups in biopolymers. It is widely used as a chemical crosslinker in the production of medical materials [45]. The concentration of glutaraldehyde up to 0.5% used in this study provides an effective degree of cross-linking without causing excessive stiffness or brittleness in the hydrogel structure [47,48].

SEPINEO™ D.E.R.M. is a polymer thickener based on hydroxyacrylate and sodium acrylodimethyltaurate. It is stable across a broad pH range (3–12) and is commonly used at concentrations of up to 3% in dermatological gels to improve spreadability and skin feel [49]. In our study, SEPINEO™ D.E.R.M. was used at a concentration of 3% in gelatin-free hydrogel formulations to achieve a smooth, non-sticky texture.

Lidocaine hypochlorite is a local anesthetic that suppresses pain signals by blocking sodium channels on the membrane of nerve cells. It is often incorporated into wound gels to provide rapid pain relief. In this study, lidocaine hypochlorite was added at 0.5%, a concentration selected to balance analgesic efficacy and safety, while avoiding risks of systemic absorption from larger wounds [50,51,52].

Chlorhexidine bigluconate is a widely used antiseptic with both bactericidal and bacteriostatic activity. It acts by disrupting the osmotic balance of microbial cell membranes. Clinically, it is effective in managing burn wounds by reducing microbial load without impeding tissue regeneration [53]. In our hydrogels, chlorhexidine was included at concentrations up to 1%, which ensured antimicrobial activity without cytotoxic effects on skin cells [54].

Chloramine T is an organic N-chloramine with antimicrobial activity that helps reduce the risk of sepsis while having low dermal toxicity. The concentration of Chloramine T we have chosen, up to 0.5%, promotes antiseptic properties while eliminating negative skin reactions [55,56].

Conditioned Dulbecco’s Modified Eagle Medium (DMEM), a waste product from stem cell culture, was used both as a solvent and as an active component in anti-burn hydrogel compositions. Its rich composition includes enzymes, proteins, cytokines, growth factors, extracellular vesicles (microparticles, apoptotic bodies and exosomes) and autophagosomes. These factors have been shown to regulate metabolic activity and promote tissue regeneration. Moreover, the conditioned medium allows for long-term storage without requiring toxic cryopreservatives such as dimethyl sulfoxide (DMSO) [57].

3.3. Physicochemical Properties of Composite Hydrogels

3.3.1. Hydrophilicity Assessment

The hydrophobicity/hydrophilicity of the 23 experimental samples was determined using a modified drop test. All studied samples dispersed in the aqueous phase and remained agglomerated in the oil phase, indicating that the hydrogels belong to the “oil-in-water” emulsion type—confirming their hydrophilic nature.

Visual assessment revealed that the water-based gels were colorless or white, while those prepared using conditioned medium displayed a pronounced color ranging from yellow to crimson. This is explained by the presence of Phenol Red indicator in the conditioned medium, which changes the color of the solution depending on the concentration of the components. All formulations exhibited a soft texture when applied to the skin. Samples No. 1, 5, 8, 12, 15, 16, 20, and 22 were found to have a sticky consistency. All tested samples showed a neutral odour and homogeneous structure.

3.3.2. pH Measurement

The pH values of the hydrogel samples are presented in

Table 3. pH values of experimental samples.

Samples No.	pH (Hydrogen Index)
1	5.37 ± 0.02
2	5.97 ± 0.06
3	8.16 ± 0.09
4	6.89 ± 0.10
5	8.64 ± 0.07
6	7.51 ± 0.08
7	6.35 ± 0.06
8	6.85 ± 0.11
9	7.12 ± 0.08
10	4.10 ± 0.07
11	7.65 ± 0.05
12	8.16 ± 0.04
13	6.70 ± 0.10
14	5.11 ± 0.08
15	4.76 ± 0.03
16	4.81 ± 0.04
17	5.29 ± 0.06
18	4.98 ± 0.11
19	4.80 ± 0.04
20	5.10 ± 0.05
21	5.40 ± 0.04
22	5.90 ± 0.08
23	5.34 ± 0.02

. The pH of topical pharmaceutical formulations is a critical parameter influencing their stability, efficacy, and skin compatibility. According to multiple sources, the optimal pH range for topical applications lies between 4 and 6 [58]. Eight hydrogel samples—No. 1, 2, 7, 14, 17, 21, 22, and 23—had pH values ranging from 5.11 ± 0.08 to 6.35 ± 0.06, thus meeting the criteria for dermal compatibility. These samples were selected for further investigation.

3.3.3. Moisture Content

An important characteristic of hydrogels is their ability to retain a significant amount of water within a three-dimensional polymeric matrix. The moisture content values of the samples studied are presented in

Table 4. Moisture content in hydrogels.

Samples No.	1	2	7	14	17	21	22	23
Moisture content, %	89	88	87	89	89	87	92	92

.

The analysis was conducted under natural evaporation at room temperature, which closely resembles the actual storage conditions of topical formulations. All samples demonstrated high moisture retention, consistent with the hydrophilic nature of the polymer matrix. According to the literature, hydrogel scaffolds with water content exceeding 90% are optimal for maintaining a moist wound environment, which facilitates healing [59]. Such moisture not only reduces pain through surface cooling but also prevents the dressing from adhering to the wound. Among the eight hydrogel samples tested, samples No. 22 and No. 23 exhibited the highest moisture content, at 92%, and were selected for further evaluation.

3.3.4. Swelling Degree

The swelling capacity of hydrogel samples was evaluated after freeze-drying. Differences in pore number and size were observed among the samples (Figure 4). Sample No. 22 displayed a surface with large pores and moderate gloss, corresponding to a swelling degree of 175%. Sample No. 23 showed a smoother, more uniform surface and achieved a swelling degree of 213%.

According to existing literature, an effective swelling degree for pharmaceutical hydrogels ranges from 100% to 300% [60]. Therefore, the high swelling capacities observed for samples No. 22 and No. 23 are likely attributable to their porous structure and high content of hydrophilic functional groups. These properties contribute to effective exudate absorption and the controlled release of active compounds to the wound site.

Differences in swelling behavior between the two samples may be attributed to variations in polymer composition. Sample No. 22, which includes chitosan, demonstrates a lower swelling degree, likely due to chitosan’s ability to form dense hydrogen and ionic bonds that increase network crosslinking and reduce water permeability [44]. In contrast, the hydrophilic polymers pullulan and gelatin, present in both samples, promote greater water retention [61].

3.3.5. Morphological Analysis

The results of the study of the morphology of lyophilized hydrogels are presented in the SEM images (Figure 5). The SEM image of hydrogel sample No. 22 shows a well-defined porous structure. The image revealed oval and spherical pore morphology with sizes ranging from 1.96 µm to 21.27 µm. The structure of the hydrogel is homogeneous, the pores have a well-defined structure with clear walls. Chitosan contributes to the formation of a branched and complex porous network [62].

Sample No. 23 has a denser structure with smaller pores than sample No. 22. The distribution of pores by size is relatively uniform. Thus, the sizes of most pores are in the range of 5.86–7.95 µm, and their maximum size does not exceed 8.70 µm. The number of pores is smaller than in sample No. 22. Both samples have through pores.

Overall, lyophilisation of the obtained hydrogels leads to the formation of a system of through pores in a wide range of sizes in both samples.

3.3.6. Dynamic Viscosity of Hydrogels

When designing modern wound-healing gel coatings, one of the key performance parameters is the rheological behavior of hydrogels, particularly their dynamic viscosity. This parameter plays a crucial role in determining the ease of application, the ability of the gel to remain on the wound surface, and its spreadability and interaction with wound exudate [5]. Dynamic viscosity was measured for the experimental samples at two temperatures—25 °C and 60 °C—and at different rotational speeds. This approach enabled assessment of the structural and mechanical behavior of the hydrogels under varying shear conditions (

Table 5. Dynamic viscosity of hydrogels.

Sample No.	Spindle	Temperature, °C	Spindle Speed, rpm	Dynamic Viscosity, mPa·s
22	#No. 62	25	30	120.0
			50	110.4
			60	91.5
			100	80.4
		60	30	18.0
			50	14.4
			60	16.0
			100	19.5
23	No. 62	25	30	109.0
			50	88.0
			60	78.5
			100	58.0
		60	30	117.0
			50	96.6
			60	85.5
			100	61.2

).

A decrease in viscosity with increasing shear rate (from 30 to 100 rpm) was observed at 25 °C in both samples, indicating pseudoplastic (shear-thinning) behavior. This response is characteristic of polymer networks partially disrupted by mechanical force, facilitating gel spreadability and application [63].

At 25 °C, sample No. 22 consistently showed higher viscosity than sample No. 23 across all tested shear rates—120 mPa·s at 30 rpm and 80.4 mPa·s at 100 rpm, compared to 109 and 58 mPa·s, respectively. This difference can be attributed to the presence of chitosan in sample No. 22, a polycationic biopolymer with high molecular weight and the capacity to form hydrogen and ionic bonds with polysaccharides [44]. The presence of chitosan contributes to enhanced internal cohesion and greater resistance to shear deformation.

Upon increasing the temperature to 60 °C, notable changes in rheological behavior were observed. Sample No. 22 showed a sharp drop in viscosity, from 120.0 to 18.0 mPa·s, indicating thermal instability. In contrast, the viscosity of sample No. 23 increased from 109.0 to 117.0 mPa·s under the same conditions. This opposing behavior likely reflects differences in thermal response of the polymer components.

Chitosan is known to be thermolabile in aqueous environments, undergoing hydrolytic degradation at temperatures above 40 °C, which leads to a reduction in molecular weight and solution viscosity [63]. The viscosity drop in sample No. 22 is consistent with this thermosensitivity.

Conversely, the thermal stability of sample No. 23 is likely due to the thermogelation behavior of gelatin and pullulan chains, which undergo structural compaction at elevated temperatures [11].

Overall, the dynamic viscosity values of both samples at room temperature fall within the optimal range for medical hydrogels—between 50 and 500 mPa·s [64,65]. These values support their suitability for topical application and confirm the formulation’s potential for wound-healing purposes.

3.3.7. Thermogravimetric Analysis

Empirical data obtained as a result of thermogravimetric analysis indicate the thermal stability of the hydrogels, which is important for biomedical applications. The analysis showed differences in the decomposition profiles of samples No. 22 and No. 23. Hydrogel No. 22 began to lose mass at temperatures T₅% = 93.4 °C and T₁₀% = 120.8 °C, while gel No. 23 at T₅% = 113.0 °C and T₁₀% = 190.9 °C. This reflects the higher hydrophilicity of sample No. 22 and the water-binding ability of chitosan in the formulation. The initial dehydration processes are explained by the release of bound water. For hydrogel No. 23, the main degradation temperature of the polymer matrix was 302 °C. Hydrogel No. 22 showed only a slight difference in thermal stability compared with sample No. 23 (T_max = 309 °C). Hydrogel No. 23 also showed a sharp single-stage degradation. The gradual loss of mass in sample No. 22 (Figure 6) suggests that the structure is stabilized by intermolecular hydrogen bonds and electrostatic interactions between polymers. The structure-stabilizing effect of chitosan has been reported in the literature [66].

The residual mass at a temperature of 600 °C was 25.1% for No. 22 and 29.3% for No. 23. This can be explained by the degree of condensation reaction during decomposition.

The described two-stage dehydration profile is typical for polysaccharide materials and hydrogels and has been confirmed by independent studies, where moisture evaporation begins at about 100 °C and lasts up to 150 °C. However, changes may occur depending on the degree of water binding by the polymer network [66,67].

Based on the results of the analysis, it can be concluded that the samples are thermally stable enough to withstand the conditions of thermal sterilization for use in medicine; however, their medical suitability can only be fully confirmed when combined with additional analyses.

3.3.8. Cytotoxicity Assessment in Cell Culture

The experimental results demonstrated that both hydrogel formulations were non-toxic, supporting cell attachment and proliferation. As shown in

Table 6. Percentage of mesenchymal stem cell viability after incubation with hydrogel samples.

Sample No.	Hydrogel Content in the Nutrient Medium, %	Cell Viability, %
		3 Days	5 Days
control	-	93 ± 2	92 ± 1
22	10	91 ± 1	91 ± 1
	30	89 ± 2	87 ± 2
	50	86 ± 3	82 ± 2
	70	76 ± 2	74 ± 1
23	10	93 ± 2	90 ± 1
	30	89 ± 1	89 ± 1
	50	88 ± 1	85 ± 2
	70	81 ± 3	78 ± 2

, cells remained viable in all tested conditions. MSC viability ranged from 91 ± 1% to 74 ± 1% for hydrogel No. 22, and from 90 ± 1% to 78 ± 2% for hydrogel No. 23.

According to GOST ISO 10993-5, samples causing a viability reduction of less than 20% are considered to exhibit negligible cytotoxicity, corresponding to the lowest level on the qualitative cytotoxicity scale [21].

3.3.9. Antimicrobial Activity of Hydrogels

An important property of wound-healing gels is their antimicrobial activity. During formulation, concentrations of antimicrobial agents were carefully selected to ensure effective topical antimicrobial action. Glutaraldehyde at a concentration of 0.1% was found to meet these criteria.

The antimicrobial activity of hydrogels No. 22 and No. 23 was assessed using the agar diffusion method. Zones of inhibition (“halo” zones) were measured against the following test microorganisms: Ps. aeruginosa 853, E. coli 603, and S. aureus ST228. These strains were selected due to their clinical relevance in the pathogenesis of burn wound infections [68,69,70,71,72].

Both hydrogels showed antimicrobial activity against S. aureus and E. coli, with inhibition zones ranging from 4.7 ± 0.1 mm to 11.3 ± 0.1 mm. Notably, only hydrogel No. 23 exhibited activity against P. aeruginosa, producing a halo zone of 4.0 ± 0.1 mm (

Table 7. Antimicrobial activity of experimental hydrogels.

Test Cultures	Growth Inhibition Zones of Test Cultures, mm
	Gel No. 22	Gel No. 23
S. aureus ST228	A 4.7 ± 0.1	B 8.8 ± 0.1
E. coli 603	C 4.8 ± 0.1	D 11.3 ± 0.1
Ps. aeruginosa 853	No activity	E 4.0 ± 0.2

).

The results of a qualitative analysis of the antimicrobial properties of the experimental hydrogels demonstrated their moderate activity against both Gram-positive (S. aureus ST228) and Gram-negative (E. coli 603, Ps. aeruginosa 853) bacteria. This activity is likely due to the presence of chitosan, glutaraldehyde, and citric acid in their composition. Chitosan exerts antimicrobial effects through disruption of bacterial membranes and leakage of intracellular contents [73]. Citric acid lowers the pH, which can be detrimental to neutrophilic bacteria that are sensitive to changes in the acidity of the environment [74]. Glutaraldehyde exhibits bactericidal activity through oxidation of cell walls [75].

A possible explanation for the reduced activity of hydrogel No. 22 against Ps. aeruginosa is its higher viscosity due to the presence of chitosan. Increased viscosity can limit the diffusion of active agents into the agar medium, reducing the apparent antimicrobial zone [76].

The activity of the hydrogel prototypes on the planktonic cells of the test microorganisms was also investigated by determining MIC values. For hydrogel No. 22 containing chitosan, complete inhibition of S. aureus ST228 growth was observed at 30% gel concentration in nutrient agar, whereas for inhibition of E. coli 603 growth, an 80% gel concentration was required. These data are consistent with studies by a number of authors emphasizing the superior activity of chitosan against Gram-positive species [77]. This susceptibility is explained by the peculiarities of the bacterial membrane: Gram-positive microorganisms have an external cell wall consisting of acidic polysaccharides (teichoic acids) and peptidoglycan, with a large number of pores, which promotes adhesion or penetration of foreign molecules into the cell [78].

Pullulan-based hydrogel No. 23, by contrast, suppressed the growth of E. coli to a greater extent. Thus, for this test culture, MIC corresponded to a 20% gel concentration, whereas a 60% gel concentration was required to completely suppress the growth of S. aureus. The hydrogels had the same effect on P. aeruginosa 853, and the MIC for this test organism corresponded to a concentration of 60% in both gels. Due to the antimicrobial nature of citric acid, polymers based on citric acid may have their own antimicrobial properties [79]. It is known that the antimicrobial effect of citric acid is associated with a local decrease in pH, which can lead to a decrease in the internal pH of bacteria and/or a change in the permeability of the microbial membrane, disrupting substrate transport [80,81,82]. Thus, unreacted carboxyl groups of citric acid-based polymers and citric acid released during degradation may be responsible for the antimicrobial effect of the hydrogel [76].

3.3.10. Inhibition of Biofilm Formation by Hydrogel Prototypes

To determine the effect of hydrogel prototypes No. 22 and No. 23 on biofilm formation, the quantitative microtiter plate assay was used. The bacteria were cultured in microtiter plates for 24 h in the presence of various concentrations of hydrogels (0.5–2.7× MIC), and the resulting biofilm was stained with crystal violet. Overall, gel No. 22 showed lower biofilm inhibition compared to gel No. 23. With 1× MIC of hydrogel No. 23, the growth of S. aureus ST228, P. aeruginosa 853 and S. aureus ST228 + P. aeruginosa 853 was almost completely inhibited (Figure 7). This result indicates that hydrogel No. 23 has an inhibitory effect on both planktonic cells and biofilm formation.

3.3.11. Wound Healing Properties of Hydrogels In Vitro

Re-epithelialization is a critical indicator of wound healing. Fibroblast migration is essential for skin restoration during all phases of wound repair. Accordingly, the final stage of this study evaluated the ability of the experimental hydrogels to promote cell migration in vitro.

Scratch assay analysis was performed to assess the effect of hydrogels No. 22 and No. 23 on fibroblast migration. The scratch assay is a simple and effective 2D model commonly used to study wound closure dynamics [83]. Observations were recorded at 3, 6, and 24 h following scratch creation and hydrogel application.

Following the scratch, fibroblasts began migrating from the intact cell layer toward the wound gap. This migration continued during incubation until intercellular contacts were re-established and the gap was closed. The process was documented at 3, 6, and 24 h using an inverted microscope, with photographs taken of marked observation areas (

Table 8. Percentage of scratch filling by fibroblasts.

Sample No.	Area of the Initial Scratch Relative to the Entire Field of View (A)	Area of the Scratch Relative to the Entire Field of View After 24 h (B)	Percentage of Scratch Filling (A–B)
22	50.27 ± 0.21%	30.87 ± 0.25%	19.40 ± 0.33%
23	52.58 ± 0.19%	22.33 ± 0.22%	30.25 ± 0.29%
control	48.49 ± 0.22%	29.26 ± 0.24%	19.23 ± 0.33%

and

Table 9. Scratch analysis results.

Sample No.	Scratches Before Application of Components	Scratches After 24 h
22	A	B
23	C	D
control	E	F

).

The results presented in Table 8 and Table 9 indicate that hydrogel No. 23, containing pullulan at a concentration of 24 mg/mL, strongly stimulated fibroblast migration. This formulation restored the adhesive properties of the damaged cells and promoted cell proliferation. The percentage of scratch closure in this variant reached 30.25 ± 0.29% after 24 h. In contrast, the sample containing chitosan (hydrogel No. 22) did not enhance fibroblast migration.

This difference is likely attributable to the biological and physicochemical properties of pullulan. As a biodegradable polysaccharide, pullulan may serve as an energy source for fibroblasts, while its hygroscopic nature contributes to the maintenance of a moist environment, which is known to facilitate cell migration [10,84].

Hydrogel No. 22, which contained chitosan, did not exhibit wound-healing effects in this assay. Several factors may contribute to this observation, including increased viscosity, altered porosity, and limited moisture transport in the gel matrix [84].

A number of studies on chitosan’s activity toward fibroblasts interpret its action in two ways [85]. Some demonstrate the stimulating effect of chitosan, others report an inhibitory effect [86,87,88]. It is believed that these discrepancies are caused by several factors, including the cell type, the method used, and the chemical properties of chitosan and its derivatives. The inhibitory effect on cell migration can also be explained by chitosan’s capacity to electrostatically bind growth factors (FGF-2), which decreases their free concentration in the medium and weakens the signals that stimulate migration [89,90,91].

In summary, the results confirm the effectiveness of pullulan in promoting fibroblast migration, highlighting its potential as a key component in hydrogels designed for wound-healing and anti-burn applications.

4. Conclusions

In this study, wound-healing hydrogels were developed and evaluated based on the microbial polysaccharide pullulan, produced by the yeast-like fungus A. pullulans C7. Structural characterization by IR spectroscopy and NMR confirmed the identity and purity of the extracted polysaccharide, consistent with literature data for pullulan composed of α-1,6-linked maltotriose units.

The developed hydrogels demonstrated several advantageous properties, including high water retention capacity, favorable rheological behavior, skin-compatible pH, and structural homogeneity. Their biocompatibility and non-toxicity were confirmed in vitro using mesenchymal stem cells, in accordance with ISO 10993-5 [21], showing no significant cytotoxicity across a range of concentrations.

Moreover, antimicrobial activity was observed against clinically relevant Gram-positive and Gram-negative bacterial strains, likely due to the synergistic action of chitosan, glutaraldehyde, and citric acid. Among the 23 formulations tested, samples No. 22 and No. 23 were selected for further investigation based on optimal physicochemical characteristics.

Importantly, scratch assay analysis demonstrated that hydrogel No. 23, which contained pullulan and gelatin, significantly promoted fibroblast migration and wound closure in vitro, thereby confirming its potential as a bioactive dressing for wound healing. In contrast, the chitosan-containing hydrogel No. 22 showed reduced cell migration, likely due to increased viscosity and limited diffusion of active compounds.

Overall, the findings highlight the importance of rational formulation and the synergistic roles of individual components in designing effective hydrogels. Pullulan-based hydrogels, in particular, show promise as multifunctional wound dressings with excellent physicochemical, antimicrobial, and regenerative properties, supporting their further development for clinical and pharmaceutical applications.