Biodegradable Foam Synthesized from Lignin-Rich Brewer’s Spent Grain Waste with Potential Use as a Sustainable Alternative for Packaging Applications

Abstract

Biodegradable foams derived from agroindustrial residues provide a sustainable alternative to existing petroleum-based packaging, while supporting circular bioeconomy objectives. In this study, biofoams were synthesized using raw brewer’s spent grain (BSG1) and lignin-enriched residues (BSG2 and BSG3) obtained postfermentation for ethanol production. Chemical composition analysis of all BSG biomasses revealed that BSG1 contained 18.72% total lignin, whereas BSG2 and BSG3 exhibited substantially higher lignin contents of 78.28 and 83.69%, respectively, representing nearly a 4-fold increase. SEM (scanning electron microscopy) analysis showed an increase in biofoam porosity from BSG1 (0.4%) to BSG2 (7.8%) and BSG3 (13.4%), which correlated with enhanced compressive strength (0.49–0.50 MPa) and improved thickness recovery under uniaxial cyclic compression. It also exhibited reduced water absorption, from 20% in BSG1 to 7 and 11% in BSG2 and BSG3, respectively. The enhanced lignin content may have contributed to the strength and moisture resistance of the biofoams, whereas the relatively low water absorption of the processed BSG likely supported its degradation in the environment. The foams exhibited high solubility in river and seawater (48–70%) and near-complete soil degradation within 7–10 weeks, highlighting their rapid environmental decomposition. Thermal analysis (TG/DTG) confirmed the multistage degradation typical of lignocellulosic materials, with higher residual char in BSG2 and BSG3 reflecting their lignin-rich composition. Collectively, these results demonstrated that lignin-rich BSG residues can be transformed into mechanically robust, moderately porous, thermally stable, and rapidly biodegradable biofoams, combining enhanced strength, controlled hydrophilicity, and environmental compatibility. This work establishes a zero-waste cascade valorization strategy in which lignin-rich fermentation residues are upcycled into biodegradable biofoams, providing a sustainable pathway for replacing petroleum-based packaging materials.

1. Introduction

In the past few decades, there has been a continuous rise in global plastic production, exceeding 370 million tons annually, which has increased the environmental burden due to the accumulation of toxic nondegradable residues in terrestrial and marine ecosystems. The extensive use of polystyrene (PS)- and polyurethane (PU)-based foams in several industries, such as packaging, pharmaceuticals, automotive, and construction, has led to large-scale irreversible damage to the environment due to their resistance to biodegradation. Expanded polystyrene (EPS) foam, commonly used for single-use packaging, is often discarded after use and persists in the environment as nondegradable waste for hundreds of years, leading to lasting environmental problems. Despite environmental concerns, EPS foams remain in high demand due to their low density, excellent thermal insulation, and mechanical robustness, supporting their extensive industrial applications. Although stringent recycling policies have been adopted globally, foam recycling remains economically and technically challenging because of its low density, contamination, and inadequate infrastructure. Consequently, regulatory frameworks, particularly within the European Union, increasingly emphasize reducing petroleum-based plastics and promoting biodegradable alternatives with targets for recyclable and sustainable packaging by 2030. To support this goal, several regulations and policies have been introduced to encourage the use of biobased and compostable materials, with the intention of replacing conventional petroleum-based plastics by 2035. , In this scenario, environmental researchers are exploring novel sources of renewable feedstocks to replace fossil-based plastics.

Biobased foams can be made available from diverse matrices like starch, polylactic acid, and cellulose-based composites, which ensure a range of biodegradation mechanisms and characteristic properties. Starch-based foams are widely used because of their abundance, renewability, and biodegradability. However, most starch-based foams tend to be highly hydrophilic and have relatively low mechanical strength, which limits their applications. , Agroindustrial byproducts that are usually rich in lignocellulosic fibers can be incorporated into starch blends to modify and enhance the characteristic properties of these existing biodegradable foams. The incorporation of sugarcane bagasse, malt bagasse, and fish scale wastes as matrix fillers has been reported to increase the density and mechanical strength and reduce the water absorption capacity of starch-based biofoams. Adopting several reinforcement and cross-linking methods can further yield biodegradable foams with enhanced porosity and cushioning characteristics, which are much-needed properties in the packaging industry.

BSG represents one such alternate solution to generate biodegradable foams. Globally, BSG is one of the most abundantly generated agroindustrial biomasses, with an estimated annual accumulation of approximately 10 million tons in Europe. Despite its substantial cellulose (15–25%), protein (20–25%), and lignin (12–24%) content, BSG remains one of the least valorized lignocellulosic biomasses, with a major fraction still being disposed of in landfills, thereby contributing to greenhouse gas emissions and environmental burdens. , The valorization of BSG for the synthesis of biodegradable foams offers a sustainable pathway to transform an underutilized organic residue into value-added biomaterials, aligning with the principles of resource efficiency and circular bioeconomy frameworks.

Recent research on sustainable materials has identified BSG as a promising feedstock for the development of biobased polymers. Through processes such as thermomolding and extrusion, BSG has been effectively incorporated into biocomposites, thin films, and lightweight foams, providing an eco-friendly alternative to conventional petrochemical products. These materials typically exhibit low bulk densities, often below 0.25 g/cm³, and undergo complete biodegradation within 3–6 months under composting conditions, aligning with the current drive toward a circular economy and sustainable materials. , However, despite its abundance and favorable composition, the potential of BSG for the synthesis of biofoams remains largely underexplored. A deeper understanding of its structural and chemical characteristics and their influence on foam formation and performance could open new avenues for the sustainable valorization of this agroindustrial residue.

In recent years, the integrated biorefinery concept has emphasized cascade biomass utilization to encourage a zero-waste approach for developing sustainable biomaterials. Lignocellulosic feedstocks from agricultural lands are sequentially utilized and converted into high-value platform chemicals, with the remaining residual fractions further upgraded into functional biomaterials, thereby focusing on efficient resource management and minimizing waste generation. BSG, a lignocellulosic-rich biomass, can be a potential candidate for the fermentation process to obtain biofuels and other platform chemicals. During this process, carbohydrate fractions are selectively consumed, yielding lignin-enriched residues that remain largely underutilized despite their inherent structural rigidity and reinforcement potential. Lignin has an aromatic, highly cross-linked structure and contains phenolic and aliphatic hydroxyl groups as well as methoxy groups, which can promote intermolecular interactions with polymer matrices. Owing to this structure, lignin-rich residues can act as reinforcing components in biofoam formulations, potentially improving their mechanical strength, thermal stability, and resistance to moisture uptake. Previous studies have demonstrated that lignin can act as an effective biobased reinforcing phase in a range of materials, including polymer composites, porous biomaterials, and porous materials for biomedical applications, owing to its aromatic structure, hydrophobicity, and ability to enhance strength, thermal stability, and moisture resistance. , In this context, the use of lignin-enriched postfermentation BSG residues as feedstock for biofoam production offers a logical route for converting an underutilized byproduct into value-added biomaterials. This approach supports cascade valorization, improves resource efficiency, and contributes to sustainable development and circular bioeconomy objectives by reducing dependence on petroleum-derived plastics.

The present study explores the potential of BSG for producing biodegradable foams, utilizing both raw BSG and lignin-enriched residues generated through CaO-assisted pretreatment and subsequent acid neutralization during ethanol fermentation. The study systematically compares these biomass streams to understand how pretreatment-induced compositional changes influence the physicochemical, mechanical, and biodegradation characteristics of the resulting foams. This study presents an integrated cascade valorization strategy for BSG, wherein the carbohydrate-rich fraction is first utilized for ethanol production, generating lignin-enriched residues that are subsequently upcycled to biodegradable biofoams. The novelty of this work lies in the direct conversion of postfermentation BSG residues into packaging-oriented foams and the systematic evaluation of the influence of compositional enrichment on their morphology, mechanical properties, thermal stability, and biodegradability. To the best of our knowledge, this is one of the first studies to demonstrate a near-zero-waste biorefinery approach for BSG, coupling bioethanol production with the valorization of the resulting lignin-rich residues into sustainable packaging materials as an alternative to petroleum-derived foams.

2. Materials and Methods

2.1. Materials

Raw BSG (hereafter referred to as BSG1) was obtained from a local brewery (Ballykilcavan Brewery Ltd., Co. Laois, Ireland). BSG2 and BSG3 were produced from raw BSG, following pretreatment and fermentation for ethanol production. For this, BSG was pretreated with CaO and subsequently neutralized with H₃PO₄ (BSG2) and H₂SO₄ (BSG3) to facilitate the release of fermentable sugars. Hereafter, the CaO + H₃PO₄ pretreated BSG will be referred to as BSG2, while the CaO + H₂SO₄ pretreated BSG will be referred to as BSG3. Collectively, BSG2 and BSG3 will be referred to as processed BSG in this study. The resulting postfermentation residues (BSG2, BSG3), along with raw BSG (BSG1), were used as feedstocks for biofoam fabrication. Potato starch (MP Biomedicals LLC, Amsterdam, The Netherlands) and 250 bloom bovine gelatin (VWR Chemicals, VWR International, Radnor, PA) were used as the primary gel-forming components for the foam formulation. Glycerol (EMPROVE-bio, Merck KGaA, Darmstadt, Germany) was used as a plasticizer to enhance the flexibility of the biofoams. Acetone (analytical grade, VWR Chemicals, VWR International, Radnor, PA) was used as the extraction solvent, and sulfuric acid (H₂SO₄, 95–98%, analytical grade, Merck KGaA, Darmstadt, Germany) was employed for lignin determination.

For biodegradation studies, river water was collected from the River Barrow (52.50298°N, 6.56021°W) on November 20, 2025, and seawater was collected from a coastal location (52.44116°N, 6.08337°W) on July 16, 2025. Commercially sourced topsoil (Westland Horticulture Ltd., Dungannon, UK) was used for soil degradation studies.

2.1.1. Extractive-Free Lignin Estimation in Raw BSG and Processed BSG

Extractive-free BSG samples were obtained using a solvent extraction method with acetone and subsequent washing with hot water to remove the solvent and water-soluble components. Acid-insoluble lignin (AIL) and acid-soluble lignin (ASL) contents were determined using the NREL laboratory analytical procedure (LAP-003); 0.2 g of moisture-free, extractive-free BSG samples were subjected to primary hydrolysis with 72% H₂SO₄ (4 mL) at 30 °C for 60 min, followed by secondary hydrolysis with dilution to 4% acid, maintained at 121 °C for 1 h. The insoluble residue was filtered, washed, dried at 105 °C, and weighed as % AIL (Klason lignin). The absorbance of the filtrate was measured at 205 nm using a spectrophotometer to determine the ASL using an absorptivity coefficient of 110 g^–1 L cm^–1. The % total lignin content was expressed as the sum of AIL and ASL.

2.1.2. Determining the Chemical Composition of BSG Biomass by Fourier Transform Infrared (FTIR) Spectroscopy

To compare the chemical compositions of both raw and processed BSG biomass, the samples were subjected to FTIR analysis. Spectra were recorded as % transmittance using a Spectrum 65 FTIR spectrometer (PerkinElmer, Massachusetts) over the range of 4000–600 cm^–1. Powder samples were placed directly onto the diamond crystal and secured with a pressure arm to ensure optimal surface contact. FTIR spectra were processed and overlaid using Microsoft Excel, and the characteristic peaks were identified and marked with dotted lines for clarity in the comparative analysis.

2.2. Synthesis of BSG Biofoams

Biofoams were prepared using both raw and processed BSG as a lignocellulosic filler in a gelatin-starch matrix. For this, 80 mL of distilled water was heated to 45 °C in a 250 mL beaker under constant stirring (250–300 rpm), and 10 g of gelatin was gradually added and allowed to bloom for 20 min to form a homogeneous solution. A mixture of 5 g of potato starch, 11.5 mL of glycerol, and 5 g of finely milled BSG in 20 mL of water was then incorporated into the gelatin solution. The temperature was increased to 70 °C and stabilized before adding 0.5 g of enzyme to catalyze partial hydrolysis for 9 min. After removing the stir bar, the viscous mixture was aerated with a hand mixer for 2–5 min to promote foaming and porosity. The mixed suspension was poured into appropriate molds and allowed to solidify at room temperature until complete gelation, and was later allowed to dry in a climatic chamber (HD-E703-100T-4, Haida, China) maintained at controlled conditions of 20 °C and 40% humidity. All the biofoams (BSG1, BSG2, BSG3) were synthesized in triplicate (a, b, c) and cut into a 1.5 cm³ size for further analysis.

2.3. Characterization of BSG Biofoams

2.3.1. Density

Three biofoam samples, along with standard commercial white styrofoam (WF) of 1.5 cm³ size, were used for density measurements. The density of the foam was determined using the ratio of the mass to the volume of the dried foam sample. The mass of each specimen was measured using an analytical balance, and the density (ρ) was calculated as the ratio of mass of the foam to volume of the foam, according to eq

$$\rho =\frac{M_{\mathrm{f}}}{V_{\mathrm{f}}}$$ 1

where ρ is the foam density in g/cm³, M _f is the mass of the foam sample in g, and V _f is the volume of the foam sample in cm³.

2.3.2. Morphological Characterization

The microstructural morphology of the synthesized biofoams was studied using scanning electron microscopy (SEM) (Thermo Quattro S ESEM, Thermo Fisher Scientific). The instrument was operated in high-vacuum mode, with an accelerating voltage of 5 kV, a spot size of 3.5, and a working distance of 15 mm to minimize electron pooling on the sample. The samples were prepared for SEM imaging by mounting each biofoam on a stub adhered to a conductive carbon tab. To expose the internal pore structure, the biofoam samples were cut upward with a razor blade at a low angle relative to the surface with uniform and dense spacing. The resulting micrographs were used for the quantitative estimation of the pore area fraction. Porosity was estimated from low-magnification SEM micrographs by using the ImageJ software. The images were converted to 8-bit grayscale and analyzed using a defined scale bar of 1 mm, and a uniform threshold was applied to distinguish the pore regions from the solid matrix. The pore area fraction was then calculated as the ratio of the total pore area to the analyzed image area and expressed as porosity (%). For each biofoam sample, multiple representative micrographs from triplicate samples were analyzed under identical conditions, and mean values were reported for comparison.

2.3.3. Colorimetric Analysis of Biofoams

The color of the biofoam samples was measured by using a PCE-CSMI colorimeter (PCE Instruments, Meschede, Germany). The device was calibrated using standard white and black references according to the manufacturer’s guidelines. For each BSG biofoam, the color parameters L* (lightness), a* (red-green), and b* (yellow-blue) were recorded from three triplicate samples. The results were then averaged to characterize the visual properties of the biofoam.

2.3.4. Moisture Content and Water Absorption Measurements of BSG Biofoams

The moisture content of the BSG biofoams was quantified using a gravimetric drying method. Twenty-four-hour-old, demolded samples of 1.5 cm³ size were initially weighed using an analytical balance to determine their initial weight (W _i). The samples were then dried in a preheated hot air oven at 105 °C for 24 h to ensure complete moisture removal. After cooling to ambient temperature, the dried samples were weighed again to obtain the dry mass (W _f). The moisture content % was calculated using eq .

$$\mathrm{Moisture}(\%)=\frac{W_{\mathrm{i}}-W_{\mathrm{f}}}{W_{\mathrm{i}}}\times 100$$ 2

Water absorption tests for the BSG biofoam samples were carried out by following a modified TAPPI T 441 om-09 method. Dried BSG biofoam samples were initially weighed (W _i) and soaked in 200 mL of distilled water at room temperature for 1 h. After 1 h, the biofoam samples were removed from the water, and the surface water was removed by blotting with tissue, followed by weighing each water-absorbed sample (W _a). Water absorption was calculated using eq

$$\mathrm{W}\mathrm{a}\mathrm{t}\mathrm{e}\mathrm{r}\mathrm{a}\mathrm{b}\mathrm{s}\mathrm{o}\mathrm{r}\mathrm{p}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n}(\%)=\frac{W_{\mathrm{a}}-W_{\mathrm{i}}}{W_{\mathrm{i}}}\times 100$$ 3

2.4. Environmental Biodegradability Assessment

2.4.1. Solubility in River and Seawater

BSG-based biofoam samples (1.5 cm³) were weighed using an analytical balance and fully submerged in 250 mL containers filled with 100 mL of each river water and seawater. Testing was conducted at a controlled temperature of 10 °C in a shaking incubator at 100 rpm to simulate flowing aquatic environments. After 7 days, the samples were retrieved with a mesh strainer, allowed to drain excess water, and subsequently oven-dried at 105 °C until a constant weight was achieved. The percentage weight loss, reflecting solubility or degradation, was calculated relative to the initial dry mass using eq .

$$\mathrm{W}\mathrm{e}\mathrm{i}\mathrm{g}\mathrm{h}\mathrm{t}\mathrm{l}\mathrm{o}\mathrm{s}\mathrm{s}(\%)=\frac{W_{\mathrm{i}}-W_{\mathrm{d}}}{W_{\mathrm{i}}}\times 100$$ 4

where W _i is the initial mass, and W _d is the final dry mass of the treated foam.

2.4.2. Degradation in Soil

For the soil burial analysis, labeled biofoams were initially weighed (W _i) and submerged at a 2 cm depth within pots filled with topsoil. The experimental pots were stored under ambient laboratory conditions for the duration of the degradation period. Weight was recorded weekly by carefully exhuming the samples, which were gently brushed off the soil particles, oven-dried, and reweighed (W _d). The percentage mass loss was calculated using eq . , The study was continued until the complete degradation of the biofoams was observed. The evaluation was carried out until the foam samples exhibited stable weights for three consecutive weeks. The mean mass loss ± standard deviation was plotted over time to characterize the biodegradation kinetics.

2.5. Compression Test to Evaluate the Mechanical Strength of the BSG Biofoam

Uniaxial compression tests were performed on cubical specimens (1.5 cm³) of biofoams BSG1, BSG2, and BSG3 using a Mecmesin TSM Pilot tester with Emperor software (Mecmesin Ltd., West Sussex, UK). The samples were subjected to a displacement-controlled cyclic protocol to evaluate the structural response, strength, and recovery of the foams. The foam samples were placed between two flat platens and compressed at a constant rate of 5 mm/s starting from 0 mm displacement. The displacement was fixed to reach a maximum value of 12 mm, which corresponded to an 80% engineering strain relative to the initial thickness (15 mm) of the foam samples. Once the crosshead bearing load reaches 12 mm target displacement, it returns to the initial starting position (0 mm) at the same rate to complete the loading–unloading cycle.

The load–displacement data were converted to stress and strain using eqs and , respectively.

$$(\mathrm{Stress})\sigma =\frac{\mathrm{load}(N)}{A_0}$$ 5

$$(\mathrm{Strain})ϵ=\frac{\mathrm{\Delta }L}{L_0}$$ 6

where ΔL is the change in length (displacement), A ₀ is the initial cross-sectional area, and L ₀ is the initial thickness.

The maximum stress was recorded as the peak value before densification, and the ultimate deformation of the sample was calculated using the last value of the displacement using eq .

$$D=\frac{d\max }{t_0}\times 100$$ 7

where D is the ultimate deformation in %, d _max is the last value of the displacement in mm, and t ₀ is the thickness of the subjected foam. Young’s modulus was determined as the slope of the initial linear region of the stress and strain curve. To define the elastic recovery or structural integrity, the thickness of the foam after the compression test was calculated based on the ultimate deformation %, which is the permanent deformation or thickness lost by the material after compression. This can be calculated using eq .

$$t_{\mathrm{f}}=t_0\times \left(1-\frac{D}{100}\right)$$ 8

where t _f is the final thickness, t ₀ is the initial thickness of the foam, and D is the ultimate deformation of the foam sample. Data was statistically correlated, and the means and standard deviations were calculated using Excel.

2.6. Characterization of Thermal Properties

Thermogravimetric analysis was performed using a Netzsch STA thermal analyzer (Netzsch STA 509 Jupiter, Germany) to assess the thermal behavior of the foam samples. Fifteen milligrams of each foam sample were placed in an alumina crucible and subjected to heat from 20 to 900 °C at a heating rate of 10 °C min^–1 under an inert atmosphere with a continuous nitrogen flow rate of 50 mL min^–1. Thermogravimetric (TG) and derivative thermogravimetric (DTG) signals were recorded to evaluate the thermal degradation behavior of the samples. Mass loss at different temperature ranges and the corresponding decomposition stages were determined from the TG–DTG data.

2.7. Statistical Analysis

The results were analyzed using one-way analysis of variance (ANOVA) at a significance level of p ≤ 0.05, followed by a post hoc Tukey’s test to identify statistically significant differences between sample groups. All statistical analyses were conducted using Minitab 21 (Minitab Inc., PA) and Microsoft Excel 365 (Microsoft Corp., Redmond, WA).

3. Results and Discussion

3.1. Estimation of Lignin from Raw and Processed BSG

The lignin content of the raw BSG1 and processed BSG (BSG2, BSG3) exhibited significant variation with a marked increase in lignin content (%) (Table ). The lignin % in raw BSG, which is a direct byproduct of the brewing industry, was determined to be 18% ± 0.48. This is in correlation with previous works, whereas the lignin for both the metal (CaO) and acid-neutralized BSG2 and BSG3 showed an enhanced lignin content of up to 78% ± 1.46 and 84% ± 0.2, respectively. These results demonstrate that selective carbohydrate solubilization from BSG using CaO pretreatment, along with acid neutralization (H₃PO₄ or H₂SO₄), liberates polysaccharides for fermentation while leaving lignin-enriched solid residues. This lignin enrichment in the fermentation residue is due to the recalcitrant property of lignin, resistant to both hydrolysis and microbial utilization processes. The enrichment of lignin in BSG2 and BSG3 suggests that acid- and metal ion-processed BSG improves the functional aromatic reinforcement in the biofoam mixture, rather than acting only as a filler. The lignin’s phenolic structure is reported to provide improved interfacial bonds within biopolymer matrices, including starch and polyurethane-based foams. This property of lignin-rich biomass can be further explored to optimize biofoams with increased structural rigidity, hydrophobicity, and thermal stability. Hence, the conversion of lignin-enriched postfermentation BSG residues (BSG2 and BSG3) into biofoams highlights the novelty of this work, as it integrates biomass pretreatment, fermentation residue upcycling, and functional foam synthesis within a single-cascade valorization framework. Such integration underscores the novelty of this first-reported process, reinforcing the circular economy framework by maximizing resource efficiency, enhancing process sustainability, and minimizing waste generation. Subsequent biofoam formulation using raw and processed BSG, along with characterization studies, provided insights into the structural and compositional effects of lignin enrichment on the foam properties.

1. Lignin Content (%) of BSG Samples Subjected to Different Pretreatment Conditions .

material	process/treatment	lignin%
BSG1	raw obtained from the distillery	18.72 ± 0.48a
BSG2	CaO-pretreated along with H3PO4 neutralization	78.28 ± 1.46b
BSG3	CaO-pretreated along with H2SO4 neutralization	83.69 ± 0.2c

^aValues are presented as mean ± standard deviation (SD) (n = 3). Different letters (a, b, c) indicate statistically significant differences among the BSG samples.

3.1.1. Chemical Composition Analysis of Raw and Processed BSG Biomass Using FTIR Analysis

The FTIR spectra of BSG1, BSG2, and BSG3 biomass revealed differences in the chemical composition of the lignocellulosic matrix. As shown in Figure , the BSG1 spectrum was relatively weak, with bands observed around 2920, 2850, 1622, 1539, 1427, 1110, and 1060 cm^–1, which are typically associated with lignocellulosic materials. Such reduced spectral intensity has been commonly reported in crude lignocellulosic biomass, whereas NaOH and other alkaline pretreatments have reportedly altered lignin–carbohydrate interactions, thereby making the associated functional groups more evident. ,

1.

Overlaid FTIR spectra of the BSG samples recorded in the range of 4000–600 cm^–1 showing the comparative spectral profiles of BSG1 (raw BSG, blue line), BSG2 (CaO-pretreated BSG neutralized with H₃PO₄, red line), and BSG3 (CaO-pretreated BSG neutralized with H₂SO₄, green line).

The BSG2 and BSG3 biomass exhibited sharper peaks with high intensity, indicating compositional differences between the raw and processed BSG. The peaks at 2920 and 2850 cm^–1 correspond to the C–H stretching vibrations, while the peaks at 1622, 1539, and 1427 cm^–1 represent the lignin aromatic vibrations along with protein amide groups, which is also a characteristic feature of lignocellulosic biomass. , The peaks observed at 1110 and 1060 cm^–1 represent the C–O–C and C–O stretching vibrations of the polysaccharide structures, which are again related to the carbohydrate content of the biomass. The peak at 668 cm^–1 represents the aromatic C–H bending of the lignin.

Compared with BSG1, the more pronounced lignin-associated bands in BSG2 and BSG3 are consistent with the lignin enrichment observed in the compositional analyses. Similar FTIR spectral changes have been reported after the chemical pretreatment of certain agro-biomasses, which were further utilized as substrates in downstream bioconversion processes. Overall, the FTIR results provide supporting evidence for the chemical differences among the BSG samples and confirm that the processed residues contain a higher relative abundance of lignin-related functional groups.

3.2. Synthesis of BSG Biofoams

Foams were synthesized in triplicate according to the formulation to evaluate consistent patterns across samples. BSG1 foams appeared lighter in color, along with surface irregularities and heterogeneous texture, which can be attributed to the unprocessed BSG having irregular cellulose and hemicellulose fibers. On the other hand, BSG2 and BSG3 foams were brighter in color, slightly toward reddish-brown tint with a very uniform surface, and BSG3 was observed to have a uniform, smoother surface among all the foams (Figure ). The visible texture and uniform surface with a darker color can be attributed to the CaO and acid neutralization process, causing lignin enrichment, leading to phenolic depositions, giving the darker color to foams made of BSG2 and BSG3. The lignin enrichment in the processed BSG residues can be attributed to the improved interfacial adhesion within the biopolymer matrix, causing a uniform and smoother surface. Such modifications to organosolv lignin and other technical lignins have been reported to enhance the surface regularity and structure of their corresponding foam systems, thereby increasing their mechanical integrity. Such lignin reinforcements have also been reported to contribute to filler-induced polymerization and support the stabilization of the cellular network of the material without the addition of synthetic surfactants. , Thus, the lignin-enriched BSGs in this study represent a promising and novel feedstock for biobased packaging systems. The study was designed to assess the effect of feedstock upgrading on biofoam performance under a fixed formulation, so that the influence of pretreatment- and fermentation-derived compositional changes could be evaluated without the confounding effect of formulation variability.

2.

Representative images of BSG-based biofoams (BSG1, BSG2, BSG3) synthesized in triplicate (n = 3 per formulation). Samples are shown from left to right: BSG1 (a, b, c), BSG2 (a, b, c), and BSG3 (a, b, c).

3.3. BSG Biofoam Characterization

3.3.1. Density

The density of the BSG biofoams was significantly higher than that of the commercial expanded styrofoam (WF) in our study. BSG1 biofoam showed a higher density of 0.86 g/cm³ than BSG2 and BSG3 biofoams with 0.54 and 0.56 g/cm³, respectively (Figure ). Although the density differences among the BSG biofoams were modest, the lower densities of BSG2 and BSG3 can be attributed to pretreatment-induced lignin enrichment. This likely enhanced the matrix interactions and promoted pore formation during the whipping stage. The incorporation of different lignins into the polyurethane foam formulation also resulted in the development of low-density and rigid foams. From Figure , it can be elucidated that all BSG biofoams are denser than the existing commercial foams, which have a reported density of 0.03–0.12 g/cm³. Further studies on the formulation can lead to developing biofoams with lower density suitable for packaging and certain commercial applications.

3.

Density of BSG-based biofoams (BSG1, BSG2, and BSG3) compared with commercial foam (WF), expressed as mass-to-volume ratio (g/cm³). Data are presented as mean ± SD (n = 3), and bars with different letters (a, b, c) indicate significant differences (p < 0.05) among the foams.

3.3.2. Scanning Electron Microscopy

The morphological features of the BSG biofoams were characterized using SEM at two different scales. The left panel (i) of Figure shows the wide-field topographical view (35× magnification,1 mm scale), and the right panel (ii) represents the high-resolution surface view (1500× magnification, 30 μm scale) for all three biofoam samples BSG1, BSG2, and BSG3, represented as A, B, and C, respectively. The low-magnification images on the left-hand side reveal a specific transition in porosity across the three BSG biofoams. Quantitative porosity analysis averaged across the sample set using 1 mm-scale images defined BSG1 as a solid matrix with negligible porosity of 0.041%. The porosities (%) of BSG2 and BSG3 increased by 8.059 and 13.78%, respectively. The high-magnification images on the right-hand side demonstrated the microstructural changes within the solid struts. The BSG1 surface was found to have textured, leaf-like fibrous structures. The biofoams BSG2 and BSG3 showed less predominant leaf-like structures, and BSG3 was also found to exhibit smoother and homogenized cell walls, which can be correlated with its highly porous nature and enhanced lignin content.

4.

Scanning electron microscopy (SEM) images of the BSG biofoams. The left panel (i) shows a wide-field topographical view (35× magnification; scale bar = 1 mm), while the right panel (ii) presents the high-resolution surface morphology (1500× magnification; scale bar = 30 μm). The samples are arranged as A: BSG1, B: BSG2, and C: BSG3, illustrating the surface structure and pore distribution of the respective biofoam matrices.

The transition in morphology observed in BSG2 and BSG3 when compared to BSG1 is primarily due to lignin enrichment by selectively breaking the lignocellulosic complex, which was facilitated by CaO and acid neutralization of the BSG slurry used for the fermentation process. Subsequent neutralization with mineral acids, such as H₂SO₄ or H₃PO₄, can further hydrolyze hemicellulose components and alter the porosity and surface chemistry of the residual biomass. This chemical shift directly impacted the material density, as porosity is inversely proportional to the density. This can be correlated with the densities determined in our study, where BSG1 showed the highest density, which can be attributed to the low pores or less internal void space, as shown in Figure A­(i). The treatment of BSG to produce fermentation residues BSG2 and BSG3 likely softened the lignocellulosic matrix, facilitating internal gas entrapment and matrix expansion during foam formation, which resulted in more porous and lower-density biofoams. The high-magnification images in Figure A­(ii) showed distinct leafing, which is a characteristic of raw untreated biomass. With increasing lignin content in BSG2 and BSG3, these fibrous features became less predominant. Lignin acts as a natural thermoplastic binder, effectively improving the rheology of the matrix, thus creating smoother homogenized cell walls, which are more prevalent seen in BSG3. Furthermore, the images preliminarily suggest cell topology. BSG1 exhibits a closed-cell structure with an irregularly packed, dense matrix and thick walls, owing to the viscoelastic nature of its high cellulosic content. In contrast, BSG2 represents a transitional state, showing an intermediate open-cell morphology, in which lignin reinforcement increases stiffness but leads to wide wall ruptures and instability during expansion. Finally, BSG3 achieves an optimal open-cell morphology characterized by uniform, robust cell walls and a stabilized matrix, suggesting that this porous architecture is key to enhancing the foam’s overall mechanical integrity. The transition from a dense, closed-cell structure (BSG1) to a lightweight, open-cell foam (BSG3) confirms that CaO and acid neutralization are the key drivers of structural modification. The pretreatment process resulted in a higher surface area and improved uniformity, which are beneficial for applications such as acoustic insulation and other packaging.

3.3.3. Color

Color measurements were performed to assess the visual appearance and uniformity of the BSG-based biofoams resulting from different treatments. Colorimetric analysis of the three biofoam samples revealed no statistically significant differences in their a* and b* coordinates. BSG1 recorded the highest a* value, indicating the strongest red tone, while its relatively low b* value placed it closer to the blue–yellow midpoint. BSG2 showed slightly lower a* values but remained within the red region, and its moderate b* value reflected a mild yellow tone. Conversely, BSG3 exhibited high positive values for both a* and b* coordinates, resulting in a dark brownish color (Figure ). The limited variation within each data set, as indicated by the small error bars, suggests that color measurements were consistent across replicates. Overall, all samples are within the reddish color range, and the degree of yellow versus blue clearly differed depending on the BSG used for the biofoam. Color development in BSG biofoams may be associated with chemical changes occurring during thermal processing, including reactions between sugars and proteins from starch and gelatin, as well as the transformation of phenolic compounds and other conjugated chromophoric structures. The darker color observed for BSG3 may further reflect stronger acid-H₂SO₄-assisted modification of the biomass during pretreatment, which could promote partial hydrolysis of carbohydrates, cleavage of lignin–carbohydrate linkages, and lignin condensation into more conjugated aromatic structures. These chemically modified residues may also have enhanced the intermolecular interactions within the gelatin–starch matrix and contributed to a more compact cell wall network, thereby influencing both color development and the improved structural integrity of the biofoam. Overall, the use of H₂SO₄-pretreated BSG3 likely resulted in yellow-brown foams due to the formation of increased conjugated chromophores.

5.

Colorimetric properties of the BSG biofoam samples derived from BSG1 (blue), BSG2 (red), and BSG3 (green). Data are presented as mean ± SD (n = 3).

Apart from the compositional aspect, the visual aesthetics and appeal related to color can also play an important role in the acceptance of bioplastics, especially for packaging applications. Many of the existing biodegradable bioplastics lack the industrial standards set for packaging materials with respect to their optical clarity and permeability. Recently, some varieties of colored bioplastics have prevailed in the market, which reflects the growing demand for visually appealing sustainable materials. , Hence, developing a naturally pigmented biofoam in our study can also be considered an advancement toward achieving the goals of sustainable circular bioeconomy .

3.3.4. Moisture Content and Water Absorption of BSG Biofoams

Moisture content and water absorption analyses were conducted to evaluate the hydrophilic behavior and water-holding capacity of the BSG-based biofoams, which are key factors influencing their stability, porosity, and potential applicability in wet environments. BSG biofoams exhibit distinct moisture and water absorption behaviors. BSG1 was found to retain the highest moisture content of 56% and gained 20% weight from water absorption within 1 h. However, the biofoams from processed BSG residues BSG2 and BSG3 displayed lower moisture contents of 40 and 41%, respectively (Figure A). A similar pattern of reduced water absorption was observed for processed BSG2 and BSG3, with only 7 and 11% increase in weight after 1 h, respectively (Figure B). These results demonstrate a clear distinction between the raw and processed BSG residues used for biofoam synthesis. The increased hydrophilicity reflects the abundance of hydrophilic sites available within the raw, unprocessed BSG1 matrix. In contrast, BSG2 and BSG3 exhibited lower moisture content and reduced water absorption. This suggests that chemical and structural modifications decrease the availability of hydrophilic sites for water binding. Similar patterns have been reported for mineral acid and alkaline pretreatment of lignocellulosic biomass, enabling the partial removal of cellulose and hemicellulose, which redistributes the lignin within the biomass, thereby reducing the accessibility of hydrophilic sites. , Lignin enrichment introduces hydrophobic aromatic regions and reduces pore connectivity, which hinders water penetration, as observed in other lignin-modified biofoam systems. , Further, acid pretreatment is also known to partially hydrolyze amorphous hemicellulose and disrupt lignin–carbohydrate linkages, thereby increasing biomass accessibility and creating localized voids. , These structural changes may have facilitated gas entrapment during the foaming process and ultimately contributed to the morphological transition into an open-cell porous structure for the BSG2 and BSG3 biofoams.

6.

Moisture content and water absorption of BSG biofoams (BSG1, BSG2, BSG3) and commercial foam (WF). (A) Equilibrium moisture content (%) measured under standard conditions; (B) water absorption (%) after 1 h of immersion. Data are presented as mean ± SD (n = 3), and bars with letters (a, b, c, d) indicate statistically significant differences (p < 0.05) among the foams.

It was observed that the commercial EPS (WF) l foam exhibited minimal moisture content (<1%) and water uptake (2.6%), which remained consistent with its hydrophobic polymer backbone and closed-cell morphology. In contrast, while BSG2 and BSG3 showed significantly lower moisture affinity and improved structural stability compared with raw BSG1, they remained considerably more hydrophilic than the commercial control. This higher moisture sensitivity highlights a fundamental performance trade-off that is common to natural biomaterials. Although high moisture retention limits the application potential of these biofoams in high-humidity or direct liquid-contact packaging scenarios, it can provide a strategic end-of-life benefit. Unlike the inert petroleum matrix of WF, the accessible hydroxyl groups within the lignocellulosic framework facilitate the necessary water swelling and accelerated microbial colonization once discarded, balancing practical commercial utility in dry-goods transport with rapid environmental degradation.

3.4. Biodegradability Studies of BSG Biofoams

3.4.1. Biofoam Solubility in River and Seawater

The solubility of biofoams was studied in both river and seawater to evaluate the disintegration ability of the biofoam matrix when exposed to a mimicked aquatic environment. Figure A illustrates the disintegration of the BSG biofoams in water compared to the commercial foams based on weight loss over a period of 7 days when immersed in river and seawater. BSG3 showed a maximum solubility of around 70% in both river and seawater, followed by BSG2 with 62% solubility in seawater and 48% solubility in river water. Both BSG2 and BSG3 showed increased solubility up to 48–50% in river water and seawater compared to BSG1, which was found to be 46 and 50% soluble in both river and seawater, respectively. The solubility of BSG foams was significantly higher than that of commercial foams, which suggests that BSG biofoams have superior solubility in water. The higher disintegration observed for BSG2 and BSG3 is likely associated with structural modifications induced during CaO and acid pretreatment, which were responsible for the alteration of the lignocellulosic matrix and weakening the intercomponent bonding within the biomass. Although the treatment is assumed to cause relative lignin enrichment, which reduces moisture uptake, the resulting structural rearrangement can also facilitate fragmentation and environmental breakdown of the biomass-derived foam. Pretreatment of other lignocellulosic biomass is also widely known to alter the structural properties of plant cell walls, leading to enhanced susceptibility of the materials to hydrolysis and environmental degradation. Similar behavior has also been reported in starch-based bioplastics and biodegradable films, where hydrophilic carbohydrate networks promote water interaction and structural disintegration. ,

7.

Environmental degradation behavior of BSG biofoams. (A) Water solubility of BSG biofoams (BSG1, BSG2, and BSG3) compared with commercial foam (WF) in river water and seawater, expressed as percentage weight loss over 7 days. (B) Soil biodegradation profile of BSG biofoams and commercial foam (WF) determined using the soil burial test, showing percentage weight change during a 10-week incubation period under natural soil conditions. Data are presented as mean ± SD (n = 3), and bars with letters (a, b, c, d) indicate statistically significant differences (p < 0.05) among the foams.

3.4.2. Soil Burial Test to Evaluate the Biodegradability of BSG Foams in Soil

The biodegradation performance of the BSG foams (BSG1, BSG2, and BSG3) along with that of commercial WF was determined by measuring the percentage difference in the weight of the foam subjected to the soil burial test. The biofoam samples were buried in the soil, and their % mass loss over a period of 10 weeks was measured. All BSG biofoams exhibited a pronounced increase in weight during the initial phase, reaching maximum values at week 2, indicating substantial moisture absorption and swelling of the biomass matrix under the soil conditions. In contrast, the commercial WF showed only a minimal increase, reflecting its hydrophobic and inert nature. After this initial phase, all BSG biofoams demonstrated a continuous decline in weight, indicating progressive material disintegration and microbial degradation in the soil. Weight loss was represented as negative values, and the percentage difference in weight was calculated relative to the initial mass of the biofoam samples. BSG1 and BSG2 showed rapid degradation, with weight loss commencing after weeks 3–4 and reaching nearly complete weight loss (100%) by week 8. BSG3 also followed a similar trend but exhibited comparatively slower degradation, retaining a stabilized weight after week 7 with a remaining residual mass of approximately 20%. This corresponded to nearly 80% weight loss by BSG3 biofoam by week 10. This observation contrasts with that of the commercial foam (WF), which exhibited only minor changes in weight throughout the study, confirming its low disintegration under natural soil conditions. The rapid degradation of BSG biofoams can be attributed to the compositional chemistry of BSG, containing biodegradable polysaccharides and proteins that serve as natural substrates for microbial metabolism in soil. , The slower degradation pattern observed for BSG3 can be correlated with its relatively higher lignin content, providing partial structural resistance despite its susceptibility to soil microbial access to slowly utilize the polysaccharides, thereby causing eventual weight loss in natural conditions. , The increased disintegration of BSG biofoams in water and soil reflects their susceptibility to hydrolytic and microbial degradation in natural environments, underscoring the eco-friendly and degradable nature of these biofoams.

3.5. Compression Test to Evaluate the Mechanical Properties of BSG Biofoams

The mechanical behavior of the BSG biofoams was evaluated based on a uniaxial compression test, which was compared to that of standard expanded styrofoam (WF). During the loading stage, all of the foams exhibited a three-stage profile with an initial linear elastic region, followed by a plateau, where the cell wall collapsed upon increasing load, and a final densification stage, where the maximum displacement was achieved and further unloading from 12 mm to 0 mm. The biofoam recovery was deduced based on the ultimate deformation experienced by the biofoam sample (Table ).

2. Mechanical Properties of BSG Biofoams Compared to Commercial EPS Foam (WF) .

sample	maximum of stress/compressive strength (MPa)	Young’s modulus (MPa)	ultimate deformation (%)	final thickness (mm)
BSG1	0.23 ± 0.01b	0.04 ± 0.005a	10.77 ± 0.81a	13.38 ± 0.07d
BSG2	0.5 ± 0.01c	0.05 ± 0.001a	18.28 ± 0.13b	12.28 ± 0.04c
BSG3	0.49 ± 0.01c	0.43 ± 0.002c	20.88 ± 0.98c	11.69 ± 0.05b
WF	0.12 ± 0.04a	0.25 ± 0.004b	41.09 ± 0.66d	8.79 ± 0.05a

^aMaximum stress (MPa), Young’s modulus (MPa), ultimate deformation (%), and final thickness (mm) measured after a 12 mm displacement cyclic compression test. Data are presented as mean ± SD (n = 3), and values with letters (a, b, c, d) indicate statistically significant differences among foams.

The mechanical robustness of the developed biofoams was evident from their absolute compressive strength values. BSG2 and BSG3 exhibited strengths of 0.50 and 0.49 MPa, respectively, which were substantially higher than those of the commercial EPS/WF control (0.12 MPa). The enhanced strength determined for BSG2 and BSG3 can be attributed to the reinforcing property of lignin within the foam matrix, which suggests the added advantage of structural modification during the treatment process. ,, Although the pretreatment supported the introduction of microvoids into the matrix, which is evident from the porosity evaluation from SEM analysis, the lignin enrichment causing scaffold formation could have compensated for the strength, enabling efficient load distribution and resistance to compression. Such typical behavior has also been reported in lignin-rich biomaterials, where a balance between matrix reinforcement and pore distribution contributes to enhanced mechanical stability. , However, this comparison should be interpreted alongside the corresponding biofoam densities, as the compressive strength of cellular materials is strongly governed by bulk density and cell wall architecture. In the present study, the processed BSG biofoams had densities of 0.54 and 0.56 g/cm³, while BSG1 showed a higher density of 0.86 g/cm³. This indicates that the pretreatment and fermentation steps could have played a role in altering the biofoam structure in a manner that improved load-bearing performance, despite the lower density of the processed residues when compared to raw BSG1. These results therefore suggest that the lignin-enriched BSG residues formed mechanically coherent and structurally stable networks under the present formulation, while future work incorporating density-normalized mechanical metrics would provide a more direct basis for comparison with low-density commercial foams.

Despite the higher compressive strength, the Young’s modulus of BSG2 biofoam remained lower than that of commercial foam WF (0.25 MPa) in our study. This behavior may be related to the viscoelastic nature of the polysaccharide protein matrix. Among the biofoams, BSG3 showed the highest modulus (0.43 MPa), suggesting that acid neutralization of the alkaline BSG slurry during the pretreatment process may have facilitated the improvement in the structural scaffold and interfacial bonding within the BSG3 biofoam network. , In addition, the presence of controlled porosity in the processed biofoams may facilitate energy absorption during compression, which contributes to the reduced permanent deformation observed in these samples.

The recovery behavior further highlights the structural stability of BSG biofoams. While WF exhibited a permanent deformation of 41.09%, BSG biofoams showed significantly lower irreversible deformations ranging from 10.77 to 20.88%. BSG3 demonstrated a favorable combination of strength, stiffness, and recovery, retaining most of its original thickness after compression.

Overall, the data indicate that pretreatment promotes the formation of a mechanically robust, lignin-enriched matrix with controlled porosity, which not only enhances compressive strength and resilience but also supports controlled moisture uptake. This combination of structural reinforcement and moderate hydrophilicity aligns with previously observed moisture content and water absorption trends and accelerates biodegradation in both aqueous and soil environments. By facilitating microbial colonization while maintaining sufficient mechanical integrity for application, BSG2 and BSG3 biofoams demonstrate strong potential as sustainable, degradable alternatives to conventional petroleum-based foams. ,,

3.6. Thermal Properties of BSG Biofoams

Thermogravimetric analysis was carried out to assess the thermal stability and decomposition behavior of BSG biofoams, providing information on the material’s performance under heat exposure. Thermogravimetric (TG) and derivative thermogravimetric (DTG) analyses of BSG1, BSG2, and BSG3 showed the typical multistage thermal degradation pattern of lignocellulosic biomass (Figure ). An initial small mass loss of 2–3% below 60 °C corresponds to the evaporation of absorbed moisture and volatile compounds, which points to the hygroscopic nature of the material. The major degradation stage is observed between 200 and 350 °C, where sharp DTG peaks represent the decomposition of hemicellulose and cellulose and the partial decomposition of lignin. , In BSG1, peaks around 269 and 320 °C correspond to hemicellulose and cellulose degradation, respectively, consistent with the reported thermal decomposition ranges of lignocellulosic polysaccharides. , BSG2 and BSG3 exhibited similar degradation regions (240–330 °C) but with broader peaks, suggesting structural modification of the biomass following CaO treatment and acid neutralization.

8.

Thermogravimetric (TG) and derivative thermogravimetric (DTG) curves of BSG-based biofoams (BSG1, BSG2, and BSG3). TG curves (solid lines) show the overall mass loss as a function of temperature, while the DTG curves (dashed lines) highlight the rate of decomposition.

At temperatures above 380 °C, a slower mass loss was observed, which can be attributed to lignin degradation and char formation. Lignin decomposes over a wide temperature range, starting from partial decomposition at 250 °C and maximum degradation at around 700 °C due to its complex aromatic structure. Accordingly, BSG2 and BSG3 showed slightly higher residual masses at 900 °C with 10 and 14%, respectively, which were higher than the residual mass of BSG1 (10%), indicating greater char formation associated with the higher lignin content in BSG2 and BSG3. Similar increases in char residue due to lignin enrichment have been reported in thermal studies of natural fibers and lignocellulosic composites. The onset of major degradation above 200 °C suggests that all BSG fractions possess adequate thermal stability for processing with common biodegradable polymers, such as starch or polylactic acid.

The thermal decomposition behavior of BSG biofoams aligns with trends reported for other biomass-derived polymers, such as thermoplastic starch (TPS) and PLA. Similar to the BSG thermal profile, the TPS materials also displayed multiple mass-loss stages, including plasticizer and starch backbone decomposition around 330 °C, while PLA typically exhibited a single sharp degradation event due to its homogeneous polyester structure. The broader degradation peaks and higher residual char in BSG2 and BSG3 reflect the influence of lignin enrichment and structural modifications from CaO and acid pretreatment, consistent with the multicomponent behavior observed in starch- and lignocellulosic-based bioplastics. This confirms that BSG biofoams possess adequate thermal stability for processing with biodegradable polymers and exhibit the characteristic lignocellulosic thermal behavior typical of eco-friendly biofoams.

4. Conclusion

This study demonstrates that BSG residues can be valorized into biodegradable biofoams with the potential to develop mechanically robust, thermally stable, and rapidly degradable materials using targeted pretreatment strategies. To the best of our knowledge, this is the first report on the production of biodegradable foams from processed BSG, which is also a postfermentation residue, demonstrating a novel cascade valorization route for the high-value utilization of this underexplored industrial byproduct. The combination of CaO pretreatment, acid neutralization, and fermentation produces lignocellulosic residual matrices that improve the foam structure, porosity, and hydrophilic balance, resulting in materials with enhanced mechanical integrity and environmental compatibility compared to conventional petroleum-based foams. The BSG biofoams degraded rapidly in both aquatic and soil environments, confirming their potential as short-lifecycle, eco-friendly materials. By transforming an otherwise low-value residue into high-performance functional biofoams, this work illustrates a cascade valorization strategy that maximizes resource efficiency and supports circular economy principles. These findings demonstrate the potential of lignin-enriched BSG biofoams as proof-of-concept for cascade valorization and their promise as sustainable alternatives for packaging and cushioning applications. They also provide a foundation for the future optimization of formulation variables, including BSG loading and lignin concentration, to better tailor the biofoam structure and performance for targeted applications.

The authors declare that the data supporting the findings of this study are available in this paper.

S.P. developed the study concept, designed the experiments, performed all experimental work, conducted the data analysis, and prepared the original draft of the manuscript. S.F. reviewed the manuscript. J.Z. contributed to the development of the study concept, assisted in drafting the manuscript, including the abstract, and provided critical revision. L.O.N. contributed to the specific experimental analyses. J.Q., J.Z., E.J.O., Y.H., and C.B. reviewed and provided feedback on the manuscript. A.C.N. developed and supervised the project, provided critical revisions, and approved the final version of the manuscript as the corresponding author. All authors have read and agreed to the final version of the manuscript.

AI-based tools (ChatGPT, Perplexity AI) were used only for language editing and improving readability. All scientific content, experimental design, data interpretation, and conclusions were generated by the authors.

The authors declare no competing financial interest.