Development and characterization of a sustainable bacterial cellulose/collagen composite membrane for enhanced wound healing and tissue regeneration applications

Abstract

This study aimed to develop a bacterial cellulose/collagen composite membrane (BC/COL) for its wound-healing ability using collagen extracted from the scales of golden grouper (Epinephelus coioides), thus providing a sustainable derived source. BC was produced by Acetobacter pasteurianus PV549389 under static fermentation, and culture conditions were optimized using response surface methodology. ANOVA analysis confirmed the significance of the model (F = 5.09, p = 0.044), identifying optimal conditions of 21.3 g/L glucose, 4.9 g/L peptone, and pH 6.7, yielding a maximum BC of 3.99 g/L. The developed BC/COL membrane exhibited a uniform morphology, enhanced thermal stability, and clear functional group interactions, confirming the successful integration of the polymer. The biological performance of BC/COL membrane was evaluated through in vitro scratch assays and in vivo excisional wound-healing studies in rats. The BC/COL membrane exhibited a safe profile toward dermal fibroblasts, significantly enhancing cell migration (91%) and accelerating wound-healing rates (3.03%/h and 1.91%/h after 24 and 48 h, respectively) compared with the control and collagen-treated groups. Histological and histomorphometric analyses revealed enhanced tissue regeneration in BC/COL-treated wounds, characterized by increased epidermal thickness (97.5 μm), elevated collagen deposition (59.4%), improved epidermal organization, and reduced inflammatory cell infiltration compared with the untreated control group. This study shows that BC/COL membrane effectively provides a sustainable and biocompatible wound dressing material, with strong potential for clinical application.

Introduction

Cellulose, a linear polymer composed of D-glucose units linked by β−1,4-glycosidic bonds that leave accessible hydroxyl groups. Cellulose is the most abundant polymer on Earth and is a key component of plant cell walls, along with hemicellulose and lignin, typically making up between 40 and 60% of their composition^1,2. In addition to its role as a pulp and paper, cellulose possesses extensive applications within and pharmaceutical industries and renewable fuel source. However, isolating cellulose from plants is challenging, requiring complex chemical processes that can irreversibly alter the plant’s structure and lead to environmental pollution³. As the global population expands, so does the demand for cellulose, necessitating increased reliance on wood and cotton resources. This escalating demand for wood and cotton, unfortunately, has detrimental effects on the global carbon cycle⁴. Bacterial cellulose (BC) is a natural biopolymer produced by specific cellulose-synthesizing bacteria, notably acetic acid bacteria (AAB) like Agrobacterium, Rhizobium, Sarcina, and Komagataeibacter, Acetobacter^5–8.

Although BC shares a chemical composition similar to plant cellulose, its distinct structural organization gives it outstanding physical attributes. Research, including a study by Lin et al.⁸, confirms that BC showed a high tensile strength, significant crystallinity, strong hydrophilicity, and excellent biocompatibility. These distinctive characteristics and adaptable nature allow for various modifications, resulting in the possibility of using BC widely in various fields. Its applications span medicine, pharmacy, food, chemistry, environmental applications, engineering, and even fashion⁹. On the other hand, Collagen is a fibrous structural protein, and it is a major component of the extracellular matrix of living organisms¹⁰. It plays a vital role in maintaining the integrity of the biological structure and the functions of various tissues, and it constitutes about 25–30% of total animal proteins¹¹. It has a wide range of applications in various industries, including the leather industry, cosmetics, pharmaceuticals (such as collagen thin films for wound healing), and the food industry. It exhibits significant properties such as high tensile strength, low antigenicity, and good biocompatibility¹². Collagen induces coagulation of blood platelets, affects cell differentiation, and contributes to wound healing¹³. Fish collagen-derived membranes are highly promising for medical applications due to their porous structure, nutrient/oxygen permeability, high cell seeding capacity, and reduced immune response. They also stimulate fibroblast and keratinocyte creation, significantly enhancing wound healing¹⁴.

The wound management market relies heavily on dressings, a segment projected for rapid expansion. The global market, valued at $7.0 billion in 2020, is forecast to exceed $11 billion by 2025. This significant growth is primarily linked to an increase in the aging population, which fuels demand for treating chronic, surgical, and sudden-onset wounds. As a result, companies worldwide are quickly expanding the array of available biopolymer-based dressings, such as hydrogels, films, and alginates¹⁵. Therefore, BC and collagen are considered biocompatible biomaterials with complementary physicochemical properties, making them attractive candidates for wound-healing applications. To address the growing interest in sustainable biomaterials for wound healing, this study aimed to optimize a culture medium derived from the Hestrin and Schramm (HS) formulation (1954) to enhance BC production. The optimized conditions were designed to establish suitable nutritional requirements for efficient BC synthesis. In addition, BC/COL biocomposites were developed and characterized to combine the mechanical strength of BC with the biological activity of collagen. The composite membranes were subsequently evaluated for their physicochemical and biological properties to explore their potential as wound-healing biomaterials. The present study introduces a novel, high-performance bioactive membrane that bridges the gap between environmental sustainability and advanced tissue engineering. While traditional bacterial cellulose (BC)/collagen composites often rely on mammalian sources, which are frequently associated with high production costs, ethical constraints, and potential immunogenic risks, this work utilizes collagen extracted from Golden Grouper scales (Table 1). Integrated within this cellulose network is the Golden Grouper-derived collagen, which retains its native triple helix conformation. While the BC offers the physical framework and superior moisture retention, the collagen provides the essential biochemical cues required for cell adhesion and proliferation. This specific combination creates a biomimetic environment that mimics the natural extracellular matrix more effectively than either component alone. By upcycling fish scale waste into a high-value collagen source and pairing it with a statistically optimized BC production strategy, we have developed a sustainable, non-immunogenic dressing that actively promotes accelerated wound closure.

Table 1.

Comparative analysis of BC/collagen-based wound dressings.

Scaffold composition	Collagen source	Key performance (cell migration/healing rate)	Sustainability/cost profile	Study/source
Composite Membrane: Bacterial Cellulose (BC) + Marine Collagen (COL).	Sustainable Marine Waste: Isolated from the discarded scales of Golden Grouper (Epinephelus coioides)	Highly Superior Kinetics: Significantly enhanced fibroblast migration by 91% and accelerated hourly wound-closure rates (3.03%/h at 24 h and 1.91%/h at 48 h).	Highly Sustainable & Optimized: Converts marine waste into high-value biomaterials. Cost-effective due to statistical optimization (RSM/ANOVA) yielding maximum BC (3.99 g/L).	Present study
Synthetic-Impregnated Membrane: Bacterial Cellulose (BC) + Polycaprolactone (PCL) + Antibiotics (Gentamicin/Streptomycin	Not Included: This study does not incorporate any type of collagen protein.	Drug-Release Dependent: Cytocompatible on BHK-21 cells; performance relies heavily on a burst-to-controlled antibiotic release to eradicate E. coli and S. aureus.	Chemically Dependent: Utilizes synthetic PCL and clinical-grade antibiotics, raising production costs and environmental impact compared to purely green biopolymers.	Das et al.16
Drug-Loaded Membrane: Bacterial Cellulose (BC) + Vaccarin (Vac)	Natural Flavonoid Glycoside: Vaccarin isolated from Vaccariae semen (soaked into the BC matrix).	Angiogenesis & Proliferation: Stimulated L929 cell proliferation in vitro (up to 137.5% at 24 h) and accelerated epithelialization via neovascularization in vivo.	Pharma-Dependent: Relies on the extraction and chemical purification of active pharmaceutical ingredients (vaccarin), increasing formulation processing steps	Qiu et al.17
Polysaccharide-Polymer Blend: Collagen Type I (COL) + Agarose (AG)	Mammalian Tissue: Type I Collagen enzymatically extracted from Bovine tendon	Ratio-Dependent Viability: Variants A (2:1) and B (1:1) maintained excellent L929 fibroblast viability (> 85%) and normal phenotype, while variant C (1:2) induced cytotoxicity.	Resource Intensive: Uses bovine-derived collagen which faces high processing costs and safety/ethical screening, blended with commercial agarose.	Gaspar et al.18
Ternary Hybrid System: BC + Collagen (COL) + Chitosan (CS).	Focuses on standard polymer blending without waste-to-wealth focus.	Hemocompatibility & Healing: Proven to be non-hemolytic, stops bleeding via red blood cell binding, and heals 2nd-degree burns.	Multi-Step Processing: Requires sequential in-situ impregnation followed by 24-hour ex-situ immersion, increasing processing time.	Pasaribu et al.19
Bioactive Phytocomposite: BC + Collagen Hydrolysate (CH) + Quercus infectoria (QI) extract.	Hydrolyzed Form: Collagen hydrolysate (CH) blended with plant-derived polyphenols (QI).	Polyphenolic Protection: High TPC (149.7–563.1 mg GAE/g) provides potent inhibition against E. coli and S. aureus; non-hemolytic.	Solvent Reliant: Requires organic solvent (ethanol) extraction for the plant galls, adding processing steps.	Adan et al.20
Multi-Component Composite: BC + Collagen + Starch + Glycerol.	Varying Blends: Incorporated collagen and plant-derived starch into fixed BC/glycerol matrices.	Composition-Driven Customization: Adjusting ratios tailored the films for different injury types; showed compact surface morphology.	Highly Hydrophilic & Soluble: High water solubility variation (12.94–69.7%) impacts stability in high-exudate wounds.	Nunes et al.15
Covalently-Bound Nanocomposite: Dialdehyde Bacterial Cellulose (DBC) + Collagen Peptide (Col-p).	Low Molecular Weight: Collagen Peptide (Col-p, MW < 1,000 Da) for deep, uniform matrix penetration.	Controlled Release & Affinity: Sustained Fickian release of peptides; triggers cross-linking with fibroblast proteins to enhance day 4–7 cell proliferation.	Chemical Modification Cost: Requires regioselective sodium periodate (NaIO4) oxidation over 24–168 h to generate aldehyde sites	Wen et al.21
Binary Hydrogel: BC + Mammalian Bovine Collagen (COL).	Mammalian Tissue: Extracted from Bovine tendon via acid-pepsin digestion.	Standard Bioactivity: Supported baseline granulation tissue formation and active angiogenesis over 30 days.	Moderate Sustainability: Utilizes mammalian tissues which face strict and costly disease-screening protocols	Moraes et al.22

Methods

Isolation of microbial isolates

To identify cellulose-producing microorganisms, we collected approximately 30 samples of fermented fruit juice from various regions across Egypt, including Borg El-Arab City, Upper Egypt, El Bahira, and Wadi El Natron. One gram of each sample was serially diluted with sterile saline solution and then plated onto sterile Modified Hestrin-Schramm’s (HS) agar medium (per liter): 20 g glucose, 5 g yeast extract, 5 g peptone, 2.7 g citric acid, and 1.17 g disodium hydrogen phosphate²³. Its pH was adjusted to 5.5, and it was supplemented with 1.0% Amphotericin before sterilization. Plates were incubated at 28 ± 1 °C for 48 h. After incubation, pure microbial isolates were obtained and individually screened for their ability to produce cellulose on HS agar. The most potent cellulose-producing bacterial isolate was then selected and preserved for subsequent research.

For further experimentation, the selected microbial isolate was prepared by first growing it in modified LB broth medium for 24 h. Its concentration was then spectrophotometrically adjusted to an optical density (OD) of 0.685. This cell suspension was subsequently diluted with sterile physiological saline to achieve a final concentration of 8.0 × 10⁸ cells/mL. We confirmed this cell count through microscopic enumeration using a Neubauer hemocytometer (Hirschmann EM Technicolour, Eberstadt, Germany). The standardized cell suspension was stored at 4 °C until it was needed for experiments.

Phenotypic and genotypic characteristics of the isolated bacterial strain

Initial identification of this bacterial strain involved Gram staining and scanning electron microscopy (SEM) imaging. This was then followed by genetic identification, which included DNA extraction and PCR amplification of the 16 S rRNA gene. The genomic DNA was extracted according to the QIAamp DNA mini kit (cat. no. 239045) supplied from QIAGEN (USA). The 16 S rRNA gene was amplified following the method described by Lagace et al.²⁴ using universal 16 S primers, where the sequence of the forward primer (F27) was (5’-AGAGTTTGATCMTGGCTCAG-3’) and the reverse primer (R1492) was (5’-TACGGYTACCTTGTTACGACTT-3’)²⁴.

The reaction was carried out in 25 µL with the following components: 12.5 µL of 2X Emerald Amp GT PCR master mix, 1 µL of 20 pmol of each primer, 6 µL of template DNA, and 4.5 µL of PCR-grade RNAase-free water. The PCR was carried out under the following conditions according to Emerald Amp GT PCR master mix (Takara) kit: Initial denaturation (15 min at 94 °C) followed by 35 cycles of denaturation (30 Sc at 94 °C) primer annealing (1 min at 56 °C), primer extension (1 min at 72 °C) and a final extension phase (10 min at 72 °C) and then cooled to 4 °C. The PCR product was analyzed using 1% agarose gel, visualized, and photographed using a gel documentation system²⁵. Purified PCR products were sequenced using a Perkin Elmer 377 DNA sequences in combination with the Dye Deoxy Terminator Cycle Sequencing Kit (Perkin Elmer, Foster City, CA)²⁶. Similarity analysis of the nucleotides was performed using the basic local alignment search tool (BLAST), and a phylogenetic tree was constructed with MEGA 11, inferred using the neighbor-joining method.

Parameters optimizing cellulose by the most potent bacterial isolate

To systematically optimize these parameters, a one-factor-at-a-time (OFAT) approach was initially employed, where each factor was varied while all others were held constant²⁷.

Static fermentation (SF)

For optimizing cellulose production, static fermentation (SF) was employed, systematically evaluating several key factors. The incubation period ranged from 0 to 14 days. Inoculum size was tested at 400, 500, 1000, 1500, and 2000 µL (from an OD = 0.685 culture), with these experiments run for 11 days at 30 °C. The medium’s initial pH was adjusted across a broad spectrum of 2.0 to 11.0 using 1 N NaOH and 1 N HCl. Various carbon sources (glucose, sucrose, mannitol, maltose, fructose, lactose, galactose, and rhamnose) were supplied at 1% (w/v), and different nitrogen sources (NaNO₃, peptone, yeast extract, casein, glycine, and ammonium sulfate) at 0.5% (w/v). Additionally, temperature effects were explored from 25 °C to 50 °C (in 5 °C intervals). The optimization of each parameter was carried out individually using a one-factor-at-a-time analysis, while keeping all other variables constant. The optimization study employed Response Surface Methodology (RSM), specifically utilizing a Box–Behnken design (BBD). This design facilitated the optimization of three key variables: carbon source concentration, nitrogen source concentration, and pH. The experimental design comprised 15 runs to effectively construct the predictive model and determine the optimal conditions. Each variable was investigated at three distinct levels: low, medium, and high, which were coded as −1, 0, and 1, respectively (as detailed in Table 2).

Table 2.

The variables used in the BBD design for optimization of BC production yield.

Run order	Independent variables			Bacterial cellulose (g/L)
	Glucose (g/L)	Peptone (g/L)	pH	Yield
1	30	5	6	2.800
2	10	2	7	1.337
3	30	2	7	1.620
4	10	8	7	1.124
5	20	5	7	4.810
6	10	5	6	1.840
7	20	8	8	0.842
8	30	8	7	1.870
9	20	2	8	1.287
10	20	2	6	2.245
11	30	5	8	0.970
12	20	5	7	3.858
13	10	5	8	0.921
14	20	8	6	1.971
15	20	5	7	2.973

Minitab software (version 18) was used to predict the optimal values for these three variables. The relationship between the variables and the dependent response was described by a second-order polynomial equation^3,28:

1

In this equation, Y represents the predicted response, specifically the Bacterial Cellulose (BC) production yield (g/L). B₀ signifies the fitted response at the design’s center point. The coefficients B_i, B_ii, and B_ij correspond to the linear, quadratic, and cross-product regression terms, respectively. X_i and X_j, represent the coded independent variables: X₁ for carbon source concentration, X₂ for nitrogen source concentration, and X₃ for pH. Before establishing the range and corresponding levels for each variable in the BBD, a preliminary one-factor-at-a-time approach was conducted, as outlined in Table 1. The complete experimental design generated by the BBD is presented in Table 3.

Table 3.

Analysis of variance.

Source	DF	Adj SS	Adj MS	F-Value	P-Value
Model	9	16.6353	1.84836	5.09	0.044
Linear	3	3.5007	1.16689	3.22	0.120
Glucose	1	0.5192	0.51918	1.43	0.285
peptone	1	0.0581	0.05814	0.16	0.705
pH	1	2.9234	2.92336	8.06	0.036
Square	3	12.8662	4.28874	11.82	0.010
glucose*glucose	1	5.0807	5.08071	14.00	0.013
peptone*peptone	1	5.4915	5.49150	15.13	0.012
pH*pH	1	4.2633	4.26329	11.75	0.019
2-Way Interaction	3	0.2684	0.08946	0.25	0.861
glucose*peptone	1	0.0536	0.05359	0.15	0.717
glucose*pH	1	0.2075	0.20748	0.57	0.484
peptone*pH	1	0.0073	0.00731	0.02	0.893
Error	5	1.8143	0.36286
Lack-of-Fit	3	0.1263	0.04209	0.05	0.982
Pure Error	2	1.6880	0.84402
Total	14	18.4496

ANOVA coefficients of the RSM model using a Box–Behnken design, X1 = Glucose concentration (g/L), X₂= peptone concentration (g/L), X₃ = pH, DF = degree of freedom, Seq. SS = sequential sum of squares, Adj. SS = adjusted sum of squares, Adj. MS = adjusted mean squares.

Detection and quantification of the produced cellulose

The production of BC was quantified after each fermentation run. Following a 5-day incubation period, the culture broth was centrifuged at 3300 × g for 20 min. For biomass determination, the resulting precipitated pellicle was combined with 90 mL of 0.1 M potassium acetate-acetate buffer (pH 5.0) and 10 mL of 20% cellulase. This mixture was then incubated at 50 °C with shaking at 100 rpm for 1 h to facilitate BC hydrolysis, as described by Kouda et al. (1998)²⁹. Subsequently, the solution was centrifuged again at 3000 × g for 20 min. The resulting precipitate was dried overnight in an oven at 80 °C before being weighed to ascertain the biomass. Regarding BC determination, the precipitated BC pellicle was treated with 0.1 N NaOH solutions at 80 °C for 30 min. This step aimed to eliminate bacterial cells and residual medium components³⁰. This NaOH treatment was repeated three times. After that, the solution was centrifuged at 3000 × g for 20 min. The isolated cellulose was then dried overnight in an oven at 80 °C and subsequently weighed. The final dry bacterial cellulose values were reported in mg per 100 mL of the initial medium.

Extraction and purification of collagen

Collagen was extracted from the Golden Groupers’ scales. Sea fish were obtained from the local fish market, Alexandria, Egypt, and transported to the laboratory on ice. For collagen extraction, the fish were euthanized by rapid chilling in an ice-water bath, followed by cervical dislocation in accordance with the guidelines for the use of fish in research, and scales were obtained. The scales were manually harvested and thoroughly rinsed under running tap water, followed by multiple washes with distilled water to remove surface impurities and mucus.

To maximize extraction efficiency and ensure the complete liberation of the organic matrix, the cleaned scales were dried to a constant weight and then mechanically pulverized into a fine powder using a laboratory-grade mill. This pulverization step was specifically implemented to significantly increase the surface-area-to-volume ratio for the subsequent demineralization and extraction steps. The resulting scale powder was stored in airtight containers at −20 °C to prevent protein degradation until further use. For extraction, 5.0 g of fish scales were initially treated with 0.1 N NaOH for 3 days to eliminate non-collagenous proteins and pigments. This was followed by washing with distilled water and drying. Subsequently, the samples underwent extraction with 0.5 M acetic acid for three days, after which the mixture was centrifuged at 10,000 rpm for 1 h at 4 °C. The resulting supernatants were pooled, and collagen was salted out by adding NaCl to a final concentration of 0.9 M. The collagen pellets were collected via centrifugation and washed three times with distilled water to ensure the removal of residual salts^31,32. The purified collagen was suspended in 0.5 M acetic acid and dialyzed using 10–14 kDa molecular weight cut-off tubing (24 Å pores) against 0.05 M phosphate buffer (pH 7.4) containing 2 mM phenylmethanesulfonyl fluoride (PMSF) at 4 °C. The dialyzed collagen was then lyophilized and ground into a fine powder²⁷.

Preparation of the BC/COL membrane

The purified BC was crushed in a multiprocessor and homogenized until a gel was formed, and it was later used in the production of the biocomposites. The formulation was prepared using cellulose and hydrolyzed collagen at an equal dry weight ratio of (1:1). The pH of the mixture was adjusted to 7.0 using 1 M NaOH to ensure collagen stability and prevent denaturation. The biocomposites were produced using the casting technique, with the formulations gradually heated to 70 °C at 60 rpm for 20 min³³. Then, 50 g of the mixture was weighed in polystyrene Petri dishes and dehydrated in a drying oven at 40 ± 2 °C under airflow for 20–24 h. To obtain a uniform thickness of 0.15 ± 0.02 mm, the sample was measured using Micrometer M372. No chemical crosslinkers were used in this study to maintain the natural biocompatibility of the components. Pure BC, collagen, and BC/COL biocomposites were stored in a desiccator containing silica gel under ambient conditions of 23 ± 2 °C and relative humidity of 60% for 10 days¹⁵. Before in vivo studies, the membranes were sterilized using UV exposure for 30 min per side.

Characterization of BC/COL membrane

The morphological, structural, and thermal properties of BC, collagen, and the BC–COL composite were thoroughly characterized using Scanning Electron Microscopy (SEM), Fourier Transform Infrared Spectroscopy (FTIR), and Thermogravimetric Analysis (TGA). SEM imaging was performed using a JEOL Model JSM IT 200 field emission instruments. Before imaging, lyophilized samples were cut into small sections, mounted on adhesive carbon stubs, and coated with a 10 nm layer of gold to enhance conductivity. FTIR spectra were recorded on an OPUS 8.5 Bruker Optik spectrometer in the range of 4000–400 cm⁻¹ using KBr pellets (10 mg of sample mixed with 100 mg of dried KBr and compressed into 10 mm discs) with a spectral resolution of 4 cm⁻¹ over 64 scans. TGA was carried out using TGA2 1152 Software version 3.20 (01709) to determine the shrinkage temperature (Ts) and shrinkage enthalpy (ΔHs) of the collagen and composite matrices³⁴. The protein composition and purity of collagen were analyzed by Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) following the Laemmli method. Laemmli et al.³⁵, using Bio-Rad molecular weight markers, and collagen-specific staining as described by Orabi et al.²⁷.

Physicochemical characterization of BC/COL membranes

The mechanical properties of the BC/COL membranes were evaluated using tensile strength 1ST (version TXST-12168-England) at a crosshead speed of 5 mm/min. The ultimate tensile strength and elongation at break were recorded for triplicate samples. The swelling ratio (SR) (1 × 1 cm) was determined by immersing dry membrane samples (W_d) in phosphate buffered saline (PBS, pH 7.4) at 37 °C. After 24 h, the excess surface water was removed, and the swollen weight (Ws) was measured³⁶. The SR was calculated using the formula:

2

The porosity was measured via the liquid displacement method using absolute ethanol³⁶. Ethanol was chosen for its superior pore penetration and non-swelling properties. First, the membrane was immersed in a graduated cylinder with a known volume of ethanol (V₁). After 10 min, the total volume of the submerged membrane and ethanol was recorded as (V₂). The membrane was then removed, and the residual ethanol volume was measured as (V₃). The porosity was calculated using the following equation:

3

Here, (V₁ - V₃) is the volume of ethanol absorbed into the pores, while (V₂ - V₃) represents the total membrane volume. All tests were performed in triplicate to ensure statistical accuracy.

Furthermore, the WVTR was determined by measuring the weight loss of water through the membrane. The fabricated membranes were tightly sealed over the openings of glass vials containing 10 mL of distilled water. The samples were placed in an incubator at 37 °C, and the weight was recorded over 24 h. The WVTR was calculated using the following formula:

4

Where is the weight loss (g), A is the exposure area (m²), and t is the time (h). The edges of the membranes were secured with Parafilm to ensure that water vapor escaped only through the porous structure of the composite.

Cell viability and cell culture

The normal human skin fibroblast (HSF) cell line, which was purchased from VACCERA and provided by the American Type Culture Collection (ATCC), was cultured in high-glucose Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% pen/strep. Cells were maintained under standard culture conditions at 37 °C in a humidified atmosphere containing 5% CO₂ and 95% relative humidity. The HSF cells were seeded at 1 × 10⁴ cells per well in 24-well plates and allowed to reach confluence. After 24 h of incubation, the tested compounds BC, collagen, and BC/COL were added at various concentrations and incubated for an additional 48 h. Then, the cells were washed and cytotoxicity was assessed using the MTT assay. Tetrazolium (0.5 mg/mL) was added and incubated for 4 hours, after which the resulting formazan crystals were dissolved in 500 µL dimethyl sulfoxide (DMSO). Absorbance was measured at 570 nm using a spectrophotometer, and the IC₅₀ values were calculated using GraphPad Prism version 8.3.0.

In vitro wound healing assay

The wound healing (scratch) assay was performed to evaluate the ability of the tested membranes to promote fibroblast migration. HSF cells were seeded at a density of 2 × 10⁴ cells per well in 24-well plates and cultured until confluence under standard conditions (37 °C, 5% CO₂). A uniform scratch was created across the cell monolayer using a sterile 200 µL pipette tip, and detached cells were removed by rinsing with (PBS, pH 7.4). The cells were then treated with BC, collagen, and BC/COL at different concentrations, while untreated wells served as negative controls. After 24 and 48 h, the cells were washed, fixed with 4% paraformaldehyde, and stained with crystal violet. Images of the wound area were captured immediately after scratching (0 h) and again after 24 and 48 h using a phase-contrast microscope at 40× magnification. The percentage of wound closure was calculated using Eq. (5), and the relative migration rate was determined using Eq. (6):

5

Where A₀ is the initial wound area and A_t is the wound area at the indicated time point.

6

In vivo wound healing study

To evaluate the in vivo wound healing effect of the BC/COL membrane, eighteen adult male albino rats (180 ± 20 g) were housed under standard laboratory conditions (22–25 °C, 12 h light/dark cycle) with free access to food and water for two weeks prior to the experiment. All experimental procedures were approved by the Ethics Committee of the Faculty of Science, Al-Azhar University, Assiut, Egypt (Approval No. AZHAR9/2025), and conducted according to institutional and national guidelines for laboratory animal care. The animals were randomly assigned into three groups (n = 6/group): untreated control, collagen-treated, and BC/COL membrane-treated groups. Anesthesia was induced using intraperitoneal pentobarbital sodium (3% w/v, 1 mg/kg). The dorsal area was shaved and disinfected sequentially using cetrimide–chlorhexidine solution (1:30) followed by 70% ethanol under aseptic conditions. A full-thickness circular excisional wound (2 × 2 cm²) was surgically created on the dorsal surface of each rat. Immediately after wound creation, the assigned topical dressing was applied to the wound bed. To ensure continuous therapy and optimal moisture management, the entire dressing system—including the active BC/COL membrane and secondary protective layers—was completely replaced with fresh materials every 2 days under sterile conditions.

To minimize infection risk, all surgical instruments and dressing materials were sterilized before use, and animals were individually housed in clean ventilated cages under controlled environmental conditions. Animals were monitored daily throughout the postoperative period for wound condition and general health status. Paracetamol (150–200 mg/kg, orally every 12 h) was administered for postoperative analgesia. Digital photographs of wounds were obtained on days 0, 3, 7, and 14. Wound closure was quantified as the percentage reduction in wound area relative to the original wound size at day 0. Wound measurements were analyzed using coded images to minimize observational bias. For histopathological evaluation, two rats from each group were sacrificed at each designated time point (days 3, 7, and 14), and full-thickness skin specimens from the wound area were collected for histological examination.

Hematoxylin and Eosin (H&E) and Masson’s trichrome (MTC) staining

Wound tissue specimens were fixed in 10% neutral buffered formalin, dehydrated through ascending ethanol concentrations, cleared with xylene, and embedded in paraffin wax. Sections were stained with hematoxylin and eosin (H&E) as well as Masson’s trichrome (MTC) to assess tissue morphology and collagen deposition, respectively³⁷. Microscopic examination of the stained sections was carried out using a light microscope. Quantitative analysis was performed using ImageJ software (National Institutes of Health, Bethesda, MD, USA). Epidermal thickness was measured on H&E-stained sections as the perpendicular distance from the basal layer to the outer epidermal surface at multiple randomly selected locations, and the mean value was calculated for each specimen. Collagen deposition was quantified from MTC-stained sections by applying a consistent color threshold to collagen-positive (blue-stained) regions and expressed as the percentage of collagen-positive area relative to the total tissue area examined. Multiple fields from different tissue sections were analyzed for each treatment group, and the mean values were used for statistical analysis.

Statistical analysis

Data were expressed as mean ± standard deviation (SD) from at least three independent experiments (n = 3), unless otherwise stated. Statistical analyses were performed using GraphPad Prism software (version 8.3.0; GraphPad Software, USA). The interactions among the tested variables and their effects on the measured responses were analyzed using one-way analysis of variance (ANOVA), followed by the least significant difference (LSD) post hoc test for multiple comparisons. Paired Student’s t-test was used where appropriate to compare two groups. The goodness-of-fit of the statistical model was evaluated using the correlation coefficient (R²), while the significance of the regression model and coefficients was assessed using the F-test and corresponding p-values obtained from ANOVA. Differences were considered statistically significant at p < 0.05. IC₅₀ values were calculated using nonlinear dose–response curve fitting analysis in GraphPad Prism. Error bars in all graphs represent standard deviation (SD).

Results and discussion

Isolation and identification of cellulose-producing bacteria

The fermented fruit juices represent a rich reservoir of microorganisms. From these sources, ten bacterial isolates were successfully recovered and purified using modified HS medium. Among these purified isolates, four demonstrated the ability to be a productive source of cellulose. The cellulose productivity of several bacterial isolates (S-2, S-3, S-4, & S-5) was evaluated when grown on modified HS medium. Among these, isolate S-3 emerged as the most potent producer of cellulose (Fig. 1A and B). Consequently, due to its superior productivity, strain S-3 was identified and selected for all subsequent investigations. Gram-negative rods stained pink/red by Gram stain were obtained (Fig. 1C). Also, the SEM image of BC producers by Acetobacter pasteurianus PV549389 is visualized in Fig. 1D.

Fig. 1.

Image of the preparation of bacterial cellulose (BC) from S-3 isolate. (A) the BC membrane in culture media; (B) purified wet BC membrane in water and sodium hydroxide. (C) Light microscopic image of Gram stain showing a red rod-shaped Gram-negative bacterium No. (S3) isolate (400X). (D) Scanning electron microscopy (SEM) image of BC producers by Acetobacter pasteurianus PV549389 (15000X).

To precisely identify the highly active cellulose-producing isolates, we amplified and sequenced their 16 S rRNA genes. These sequences were then submitted to the NCBI GenBank, receiving Accession Numbers PV549389. The 16 S rRNA gene PCR amplification, performed with F and R primers, consistently yielded a fragment of approximately 1500 base pairs (bp). The obtained 16 S rRNA sequences were compared against the NCBI Nucleotide Database using the Basic Local Alignment Search Tool (BLAST) algorithm. While initial partial alignments showed some degree of similarity to various genera, including Bacillus, a comprehensive phylogenetic analysis was conducted to resolve the isolate’s identity. Using the Muscle program (ClustalW2 alignment format) for multiple sequence alignment, the sequence of producing strain S-3 was compared against a curated set of reference genomic databases. This rigorous analysis confirmed a high sequence similarity of 98% with Acetobacter pasteurianus. The identification was further corroborated by the isolate’s Gram-negative morphology and its characteristic ability to produce a robust cellulose pellicle under acidic culture conditions, definitively distinguishing it from the Bacillus genus (Fig. 2). Based on this comprehensive 16 S rRNA gene sequence analysis and observed phenotypic characteristics, the isolate S-3 was conclusively identified as Acetobacter pasteurianus and formally designated as Acetobacter pasteurianus PV549389. Lin et al. (2016a, b) identified Komagataeibacter intermedius FST213-1, isolated from fermented fruit juice, as a BC producer through 16 S rDNA sequencing and biochemical tests³⁸.

Fig. 2.

Phylogenetic analysis of the position of Acetobacter pasteurianus PV549389.

Cellulose production optimization

Figure 3A shows that BC productivity increased progressively with the incubation period, reaching its peak at day 11. Productivity then remained stable until day 13, after which a decline was observed. Thus, longer culture duration generally promoted greater BC accumulation. As illustrated in Fig. 3B, the optimal pH range for BC production was between 6.5 and 7.5, with a noticeable decrease in cellulose yield when the pH dropped below 6.5. Figure 3C indicates that the optimal temperature for maximum cellulose production by the bacterial isolate was 30 °C. As shown in Fig. 3D, glucose was identified as the most effective carbon source, yielding the highest BC yield. In comparison, BC production decreased successively when using sugarcane, fructose, lactose, sucrose, maltose, glycerol, and mannitol. Furthermore, as presented in Fig. 3E, peptone used as the sole nitrogen source resulted in the highest BC yield by Acetobacter pasteurianus PV549389, outperforming yeast extract, urea, ammonium sulfate, and sodium nitrate. Notably, no BC formation was detected when asparagine or glutamine served as nitrogen sources.

Fig. 3.

Effect of different parameters on BC productivity obtained by Acetobacter pasteurianus PV549389 under SMF (A) Effect of incubation period; (B) Effect of different pH; (C) Effect of different incubation temperatures; (D) Effect of different Carbon; (E) Effect of different Nitrogen.

Response surface methodology (RSM) optimization

We selected glucose and peptone to culture Acetobacter pasteurianus PV549389 for BC production. To find the ideal concentrations of these nutrients and the optimal culture pH, we used RSM, specifically focusing on three variables: glucose concentration (X1), peptone concentration (X2), and pH (X3). First, we conducted individual experiments for each variable to understand their impact on BC yield using a one-factor-at-a-time approach. We tested glucose concentrations (X1) from 0.0 to 30 g/L. We found that 20 g/L of glucose yielded the highest BC. Interestingly, Acetobacter pasteurianus PV549389 still produced a small amount of BC even without added glucose, suggesting it might use peptone and citric acid as alternative carbon sources. For peptone (X2), we tested concentrations from 0 to 8 g/L. A peptone concentration of 5 g/L significantly increased BC yield. Importantly, Acetobacter pasteurianus PV549389 failed to produce BC in the absence of peptone, highlighting the essential role of a nitrogen source for BC production. The BC production was also observed under a wide range of pH (X3) values, from 4.5 to 8.5, with the highest yield obtained at a pH of 7. There is very low BC synthesis observed at pH levels of 4.5 or above 8.5. The RSM model was designed according to the results of the individual experiment for each variable. The data set for RSM is summarized in Table 3, and the corresponding fitting between experimental and theoretical data is given in Table 4. The second polynomial order model obtained for BC yield (Y, g/L) is as follows:

Table 4.

Model summary.

S	R-sq	R-sq(adj)	R-sq(pred)
0.602379	90.17%	72.47%	68.46%

The R² value of 90.17% was adequately high, indicating that the model explains a substantial proportion of the variability in the BC yield and demonstrating its overall significance. The optimized values for each variable as predicted by the model are X1 = 21.3 g/L, X2 = 4.9 g/L, and X3 = 6.7.

Based on the predicted model, the BC yield (Y) was found to be 4.135 g/L. A validation experiment using the optimized culture medium yielded a BC yield of 3.988 g/L. The difference between the predicted and experimental values is only 3.55%, which confirms the excellent correlation of the obtained RSM model with the experimental data. ANOVA for the RSM model revealed its overall significance (P-value = 0.044, F-value = 5.09). Within the model, all quadratic terms (glucose*glucose, peptone*peptone, and pH*pH) were found to be statistically significant (P-values of 0.013, 0.012, and 0.019, respectively), indicating a strong non-linear relationship and the presence of optimal points for these factors. Conversely, no significant two-way interaction terms were observed (P-value for 2-Way Interaction = 0.861), suggesting that the effects of the factors on BC yield are largely independent of each other within the studied ranges. The three-dimensional relationships between the tested variables and BC yield (g/L) are illustrated in the contour plots (Fig. 4A–C), which clearly delineate the optimal regions for each pair of factors. The plots indicate that specific concentrations of glucose, peptone, and pH levels lead to maximum BC production. The highest yield (> 3.5 g/L, shown in the darkest green regions) was achieved when glucose ranged between 20 and 25 g/L, peptone between 4 and 6 g/L, and pH between 6.5 and 7.5. The concentric pattern of the contour lines reflects the significant quadratic effects of all three factors, as supported by ANOVA results (p < 0.05 for glucose², peptone², and pH²). These findings confirm that BC yield follows a curved response to changes in the independent variables, justifying the application of RSM for optimization. Overall, BC production was favored under near-neutral to slightly acidic conditions (pH 6.5–7.5).

Fig. 4.

Contour plot of BC yield showing the interaction between the three tested variables. (A) peptone and glucose concentration (g/L); (B) pH and glucose concentration (g/L); and (C) pH and peptone concentration.

Extended fermentation in BC production can reduce efficiency and overall yield^8,39, with a 6-day culture specifically yielding the highest BC production at 1.023 g/L³. The reduction in pH during fermentation is primarily due to the accumulation of organic acids like gluconic, acetic, or lactic acids. Research consistently highlights the strain-specific nature of optimal carbon source utilization for BC synthesis; for instance, fructose is ideal for A. xylinum (ATCC 700178) over other sugars^38,40, while mannitol is best for G. xylinus strains^41,42. Nitrogen source also plays a critical role, with peptone supporting 0.86 g/L yield⁴³ and casein hydrolyzate yielding 4.8 g/L (Matsuoka⁴⁴ et al., 1996). In contrast, asparagine and glutamine proved ineffective for some Acetobacter sp.⁴⁵, underscoring strain-specific nitrogen requirements. When glucose is the carbon source, maintaining a more alkaline pH helps minimize gluconic acid formation, which otherwise inhibits BC production^46,47. This differs from fructose, where minimal gluconic acid forms even at low pH. Although many BC-producing bacteria prefer slightly acidic conditions^48,49, a higher starting pH can help maintain an optimal range as gluconic acid accumulates⁵⁰. The response surface methodology, specifically using a Box–Behnken design, successfully investigated how fructose concentration, peptone concentration, and pH interact to influence the BC production yield of Komagataeibacter intermedius³. Barud et al. (2008, 2011) have contributed observations on the characterization of pure cellulose.

Extraction and purification of collagen

Collagen was successfully acid-solubilized and extracted from the scales of Golden Groupers, achieving an approximate 40% yield (w/w). The purity and structural integrity of the extracted collagen were confirmed using SDS-PAGE, employing rat tail tendon collagen as a reference. All fish collagen samples exhibited consistent electrophoretic patterns, revealing characteristic protein bands at 120, 130, 200, and 250 kDa (Fig. 5). This extracted collagen consists of γ chains trimers, β-chains dimers and two α chains (α1 and α2) components.

Fig. 5.

SDS-PAGE (10%) profile of collagen isolated from Golden Groupers. Lane 1 represents the pre-stained protein molecular weight marker (kDa); Lane 2 displays the purified collagen fraction.

Characterization of BC, collagen, and BC–COL composite membrane

The SEM image of BC by Acetobacter pasteurianus PV549389 is visualized in Fig. 6A. The surface of BC reveals a compact, uniform, and continuous cellulose matrix composed of an interwoven network of elongated cellulose microfibrils exhibiting a high aspect ratio⁵¹. The SEM images confirmed that the native, freeze-dried collagen formed homogenous, multi-layered aggregates exhibiting a rigorous and highly fibrillary structure (Fig. 6B)^32,52. Following treatment of BC with the collagen solution, noticeable morphological changes were observed. As shown in Fig. 6C, the surface SEM images of the BC/COL composites reveal that the BC fibers remain visible on the surface. However, a smoother layer appears across most areas, resulting from the collagen coating that partially covers the BC network.

Fig. 6.

SEM image of (A) Bacterial cellulose (BC) produced by Acetobacter pasteurianus PV549389 cultured in a HS medium; (B) Collagen; (C) BC and collagen membrane.

The chemical structures of the BC, COL, and the resulting BC/COL composite were evaluated via ATR-FTIR to confirm successful integration (Fig. 7). The spectra for BC (Fig. 7A) exhibited the characteristic fingerprints of Cellulose I, marked by a broad -OH stretching band at ~ 3342 cm⁻¹ and the beta-glycosidic linkages at ~ 896 cm⁻¹. The observed FTIR results agreed with the peaks characteristic of cellulose I^1,53. The collagen spectrum (Fig. 7B) confirmed the preservation of its protein secondary structure, with the Amide I (C = O stretching) and Amide II (N-H bending) bands appearing prominently at 1662 cm⁻¹ and 1553 cm⁻¹, respectively. The presence of the Amide III band at 1241 cm⁻¹ serves as a definitive marker for the structural integrity of the collagen triple helix. The provided FTIR spectra similarity to collagen isolated from other species indicates a consistent molecular structure across different sources^32,52. In the BC/COL composite spectrum (Fig. 7C), the simultaneous presence of both cellulosic glycosidic peaks and protein amide bands confirms the successful formation of the bio-composite. Notably, a slight shift in the Amide I and Amide II peaks, combined with a broadening of the -OH stretching region (3300–3500 cm⁻¹, indicates the formation of intermolecular hydrogen bonds between the hydroxyl groups of the BC and the amide groups of the collagen. This molecular synergy suggests that the collagen is not merely trapped but is chemically integrated within the BC network, facilitating the mechanical stability and bioactivity required for wound healing applications.

Fig. 7.

(A) ATR-FTIR spectra of BC produced by Acetobacter pasteurianus PV549389. (B) ATR-FTIR spectra of collagen isolated from Golden Groupers’ scales. (C) ATR-FTIR spectra of BC/COL membrane.

The TGA analysis of the BC membrane (Fig. 8A) revealed two main degradation stages. An initial mass loss of 5.8% occurred between 40 and 150 °C due to the evaporation of volatile compounds and adsorbed water. The primary decomposition took place rapidly between 300 and 400 °C (peaking at 360 °C), resulting in a 77.7% mass loss attributed to cellulose pyrolysis. This profile, yielding 16.5% residual char, is characteristic of highly crystalline cellulose I. Barud et al.2008 and 2011 have contributed observations regarding the characterization of pure cellulose^51,54. In comparison, the TGA analysis of the collagen sample (Fig. 8B) shows two primary stages of thermal behavior³². The first stage, occurring up to approximately 140 °C, represents a 15.6% mass loss attributed to the evaporation of absorbed and bound water, confirming the biopolymer’s high hydrophilicity. The second major event is the thermal degradation of the organic protein structure, which begins after 140 °C and continues up to 580 °C. This stage involves the decomposition of the peptide backbone, with the TGA showing an intermediate loss of 30.8% up to ∼350 °C.

Fig. 8.

TGA thermograms showing the thermal stability and decomposition profiles of (A) TGA curve of BC produced by Acetobacter pasteurianus PV549389; (B) TGA curve of collagen isolated from Golden Groupers’ scales.

Notably, when integrated into the BC network, the composite benefits from the BC’s superior thermal resistance. The improved thermal stability may result from interactions between collagen and the crystalline BC network. This improved thermal profile is essential for ensuring the structural integrity of the wound dressing during storage and clinical application.

Mechanical and physical performance of BC/COL membranes

The BC/COL composite membrane exhibited balanced physicochemical properties suitable for wound care applications. The tensile strength was found to be 17 ± 1.1 MPa with an elongation at break of 18.2 ± 1.4%, indicating that the incorporation of fish collagen into the BC network maintains sufficient mechanical durability for clinical handling. The swelling capacity reached 370 ± 12% within 24 h, suggesting a high capacity for exudate absorption. This is attributed to the hydrophilic nature of both the BC fibers and the collagen triple helix. The porosity was recorded at 82 ± 2.4%, providing an interconnected pore structure essential for gas exchange. Notably, the WVTR was measured at 2300 ± 84 g/m²/day. This value falls within the optimal range for maintaining a moist wound environment, which prevents dehydration of the wound bed while simultaneously preventing the accumulation of excess fluid that could lead to skin maceration. The high porosity and optimal WVTR are directly correlated to the optimized 3D-fibrillar architecture of the BC, which remained uncollapsed after the integration of the Golden Grouper collagen. This structural integrity is critical for supporting the cellular infiltration observed in the subsequent in vivo studies. Furthermore, the utilization of Golden Grouper-derived collagen not only provides a sustainable, non-immunogenic alternative to mammalian sources but also demonstrates a superior ability to integrate into the BC network without disrupting the essential 3D-fibrillar architecture. This synergy between the high-purity microbial cellulose and the marine collagen triple helix results in a bio-mimetic scaffold that effectively replicates the mechanical and biological microenvironment of the native extracellular matrix.

Cell viability and cell wound healing

The MTT assay results (Fig. 9A) demonstrate that incorporating bacterial cellulose into the collagen matrix enhances the cytocompatibility of the resulting composite. BC-treated HSF cells maintained normal morphology across all tested concentrations (0–500 µg/mL), showing no detectable cytotoxic effects. In contrast, collagen alone exhibited mild toxicity at higher doses, reflected by an IC₅₀ of 339.01 µg/mL. Interestingly, the BC/COL composite showed improved cell viability, with an IC₅₀ of 669.52 µg/mL. The dose–response curves (Fig. 9B) and calculated IC₅₀ values (Fig. 9C) confirm the superior biocompatibility of BC/COL compared to collagen alone. This enhancement may be due to the inherently biocompatible nature of BC and its ability to regulate the release of collagen into the culture medium, reducing localized concentration spikes and improving fibroblast tolerance⁵⁵.

Fig. 9.

(A) Bright-field microscopic images of HSF cells showing normal cellular morphology and integrity at non-cytotoxic concentrations of the tested compounds. (B) Dose–response curves of HSF cell proliferation following exposure to various concentrations of BC, collagen, and BC/COL for 48 h. (C) IC₅₀ values (µg/mL) of the tested compounds. Data are presented as mean ± SD (n = 3). Statistical analysis was performed using two-way ANOVA followed by the appropriate post hoc test (***p < 0.001 and ****p < 0.0001). Representative images were acquired using a light microscope (100× total magnification; scale bar = 100 μm). To evaluate the in vitro wound-healing potential, the scratch assay demonstrated that the BC/COL composite significantly accelerated scratch closure compared with the control group (Fig. 10A). The rate of wound closure increased in a dose-dependent manner, where higher collagen-loading concentrations (80 µg/mL), markedly enhanced HSF wound closure after 24 and 48 h (Fig. 10B and C). Since scratch assays may reflect the combined effects of both cell migration and proliferation, the observed enhancement cannot be attributed exclusively to migration. Nevertheless, the improved closure response may be associated with the synergistic interaction between BC and collagen, where the porous and biocompatible BC scaffold supports cell adhesion and spreading, while collagen provides biological cues that promote cellular activity during tissue repair. Previous studies have demonstrated that collagen plays a critical role in maintaining tissue structural integrity, regulating extracellular matrix remodeling, and supporting epithelialization during the later stages of wound healing⁵⁶.

Fig. 10.

(A) Bright microscopic images showing the migration of human skin fibroblasts (HSF) at 0, 24, and 48 h following treatment with collagen and BC/COL membranes at concentrations of 40 and 80 µg/mL. (B) Quantitative analysis of wound closure percentage (%). (C) Relative migration rate (%/h) measured after 24 and 48 h. Data are presented as mean ± SD (n = 3). Statistical analysis was performed using two-way ANOVA followed by the appropriate post hoc test (*p < 0.05, **p < 0.01, and ***p < 0.001). Representative images were acquired using a light microscope (40× total magnification; scale bar = 200 μm).

In vivo wound healing and histological assessment

Visual examination of the wounds in tested animals revealed a moist and healthy appearance, with no visible signs of inflammation or exudation from the wound edges. To evaluate the in vivo wound healing efficacy of the membrane, a full-thickness skin defect rat model was employed. Representative macroscopic wound images contraction across different treatment groups—including control—are presented in Fig. 11 (days 3, 7, and 14). Quantitative evaluation revealed that after seven days, wound closure rates were comparable among all groups, with no significant differences. By day 14, however, the wound contraction rate in the BC-COL-treated group reached approximately 64% (Fig. 12), which was significantly higher than the other groups (p < 0.01), suggesting a more pronounced healing effect.

Fig. 11.

In vivo images of wound healing using control, collagen, and BC/COL composite membrane after treatment for different periods.

Fig. 12.

Percentage of wound closure measured on days 3, 7, and 14 relative to the initial wound area at day 0 for each experimental group. Six rats were included per experimental group (n = 6), while quantitative image analysis was performed using three representative measurements per group. Statistical analysis was performed using two-way ANOVA followed by the appropriate post hoc test (**p < 0.01, ***p < 0.001, ****p < 0.0001).

Hematoxylin and eosin staining in Fig. 13A, showing a time-dependent difference in epidermal regeneration, inflammatory resolution, and dermal organization across treatment groups. At day 3, saline-treated wounds exhibited disrupted epidermal continuity, prominent crust, and abundant inflammatory infiltrates, consistent with an early inflammatory stage and delayed repair. By contrast, wounds treated with the BC/COL membrane showed early signs of epithelial bridging and developing granulation tissue. By day 7, the BC/COL group demonstrated complete re-epithelialization with a continuous stratified squamous layer and a well-formed dermal architecture. The regenerated epidermis was supported by numerous vertically oriented capillaries and an organized granulation matrix, indicating angiogenesis and progression into the proliferative phase. In the control group, epithelial defects and marked inflammatory cell presence persisted at this time point. At day 14, tissues from the BC-COL-treated animals displayed mature skin architecture with fully restored epidermal morphology, visible cutaneous appendage neoformation (hair follicles and sebaceous structures), and dense collagen deposition, and minimal inflammatory infiltrate—hallmarks of advanced remodeling. In contrast, control wounds continued to show incomplete epithelialization, scattered necrotic debris, and limited collagen formation. Masson’s trichrome staining (Fig. 13B) corroborated these observations by specifically highlighting extracellular matrix maturation. Collagen fibers (blue) were minimal in control wounds at early time points, whereas treated groups exhibited progressively greater collagen area and fiber organization. The BC/COL membrane induced the most pronounced collagen deposition and alignment by day 14, indicating accelerated matrix synthesis and remodeling relative to all other groups. Quantitative ImageJ analysis demonstrated a time-dependent increase in both epidermal thickness (Fig. 13C) and collagen deposition (Fig. 13D) in all experimental groups. Epidermal thickness increased from 28.1, 35.7, and 55.0 μm at day 3 to 54.7, 66.7, and 97.5 μm at day 14 in the control, collagen, and BC/COL groups, respectively. Notably, the BC/COL-treated wounds exhibited significantly greater epidermal thickness than both the control and collagen-treated groups throughout the healing period, reaching approximately 97.5 μm at day 14, which was 1.8-fold higher than the untreated control group. Similarly, collagen deposition progressively increased over time in all groups, rising from 13.1%, 16.1%, and 18.9% at day 3 to 42.8%, 56.5%, and 62.0% at day 14 in the control, collagen, and BC/COL groups, respectively. BC/COL increased collagen deposition by approximately 34.4% compared with the untreated control group at day 14.

Fig. 13.

Histological and histomorphometric evaluation of wound healing in different treatment groups at 3, 7, and 14 days. (A) Representative H&E-stained rat skin sections showing wound healing progression, including re-epithelialization of the wound surface (red arrows), hair follicles and neoformation of cutaneous appendages (black arrows), and inflammatory cell infiltration (green arrows). (B) Representative Masson’s trichrome-stained (MTC) sections demonstrating collagen deposition and tissue remodeling. In MTC-stained sections, collagen fibers appear blue, cytoplasmic and muscle components appear red, and nuclei appear dark blue. Increased blue staining intensity indicates enhanced collagen deposition within the wound area. (C) Quantitative analysis of epidermal thickness (µm) measured from H&E-stained sections. (D) Quantitative analysis of collagen deposition (%) determined from MTC-stained sections expressed as the percentage of collagen-positive area relative to the total tissue area. Data are presented as mean ± SD (n = 3). Statistical analysis was performed using two-way ANOVA followed by the appropriate post hoc test (*p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001). Representative images were acquired using a light microscope (40× total magnification; scale bar = 200 μm).

The combination of cellulose and collagen scaffolds represents a promising biomaterial for wound dressing applications⁵⁷. Integrating both components has been shown to enhance the in vitro proliferation of fibroblasts. Collagen plays a vital role in maintaining biological and structural support similar to that of the native extracellular matrix (ECM). It forms a dynamic system where continuous remodeling of its components regulates cellular activity and tissue functionality⁵⁸. In addition, collagen exhibits surface-active properties that allow it to interact with lipid-free interfaces. When a collagen-based scaffold is applied to an ulcerated site, it facilitates fibroblast recruitment to the wound area, promoting tissue repair⁵⁹. Conversely, cellulose contributes by absorbing exudates, maintaining a moist environment, and supporting granulation tissue formation. Together, the dual functionality of cellulose and collagen in the composite dressing enhances ulcer healing by simultaneously encouraging fibroblast activity and managing wound exudate⁶⁰.

Conclusion

The increasing need for biocompatible and sustainable wound-dressing materials encouraged the development of the BC/COL membrane in the present study. Acetobacter pasteurianus PV549389 successfully produced bacterial cellulose under optimized fermentation conditions using Box–Behnken design. The incorporation of collagen into the BC matrix improved the membrane characteristics and supported its potential application in wound healing. In vitro and in vivo findings demonstrated enhanced wound closure and tissue repair in the BC/COL-treated group compared with the untreated wounds. Histological examination also showed improved re-epithelialization, collagen deposition, and lower inflammatory infiltration in the treated tissues. Overall, the developed BC/COL membrane showed promising wound-healing activity and may represent a suitable candidate for further wound-dressing applications. However, this study was limited to a preliminary biological evaluation and did not investigate the detailed molecular mechanisms involved in the healing process. In addition, comparisons with commercial wound dressings and infected wound models were not included. Therefore, further studies are needed to evaluate the performance of the developed membrane under more clinically relevant conditions and to better understand its mechanism of action during wound healing.

Author contributions

H.A.: Methodology, investigation, formal analysis, writing—original draft, review and editing, software, data curation, and validation. M.S.: Methodology, formal analysis, data curation, writing—review, validation, supervision, and editing of the final manuscript. E.N.H.: Methodology, writing—review, validation, supervision, and editing of the final manuscript. M.Z.: Methodology, writing-reviewing, validation, supervision, and editing of the final manuscript. Y.A.E.: formal analysis, Methodology, writing—original draft, review, software, data curation, validation, and editing of the final manuscript. E.M.E.: Conceptualization, formal analysis, methodology, writing—original draft, review and editing, software, data curation, visualization, supervision, and validation. All authors have reviewed the manuscript.

Funding

Open access funding provided by The Science, Technology & Innovation Funding Authority (STDF) in cooperation with The Egyptian Knowledge Bank (EKB).

Data availability

All data generated or analyzed during this study are included in this published article.

Declarations

Competing interests

The authors declare no competing interests.

Ethical approval

Animal experiments were conducted in compliance with established standards, and all procedures were performed in compliance with applicable rules and regulations. The Faculty of Science at Al-Azhar University in Assuit, Egypt, created institutional rules for the care and use of research animals, and all animals were treated humanely in compliance with these recommendations (approval number: AZHAR9/2025).