Kraft Lignin as an Antimicrobial Agent: Effects of Solubilization Strategies on Antimicrobial Activity and Potential Biomedical Application

Abstract

The rise in microbial resistance has driven the search for sustainable alternatives, such as kraft lignin, an abundant biopolymer with antimicrobial potential. This study evaluated four methods of lignin solubilization (magnetic stirring with ultrasound, decoction in a rotary evaporator, decoction with ultrasound, and ethanolic extraction by Soxhlet) and their influence on the physicochemical, structural, and biological properties of the extracts obtained. The samples were characterized in terms of pH, solids content, molecular weight, polydispersity, morphology, functional composition, and phenolic hydroxyl content, as well as antimicrobial activity against Candida albicans and Staphylococcus aureus, and cytotoxicity. FTIR analysis indicated the preservation of syringyl and guaiacyl structural units in all extracts. Soxhlet extraction with ethanol yielded the highest molecular weight (1103 g/mol) and phenolic hydroxyl content (3.253 mmol/g), which correlated with greater antifungal activity (MICs of 0.625 and 0.312 mg/mL). Decoction in a rotary evaporator showed lower activity against C. albicans, but demonstrated antibacterial action against S. aureus (MIC 0.625 and MBC 2.5 mg/mL). The extracts were considered non-toxic, and both showed significant cell viability. The results demonstrate that solubilization methods directly influence the bioactivity of kraft lignin, highlighting its potential as a bioactive agent in sustainable microbiological control strategies.

Keywords:
kraft lignin; antimicrobial activity; Candida albicans; Staphylococcus aureus.

HIGHLIGHTS

• Kraft lignin shows sustainable antimicrobial potential.

• Solubilization strategies alter lignin structure and bioactivity.

• Soxhlet ethanolic extraction boosts phenolic content and antifungal action.

• The extracts are biocompatible, non-toxic, and maintain cell viability.

GRAPHICAL ABSTRACT

INTRODUCTION

Lignin is one of the primary components of lignocellulosic biomass, accounting for between 15% and 30% of its composition, depending on the plant source [¹]. It is an aromatic biopolymer widely available as a byproduct of the pulp and paper industry, which generates millions of tons of discarded annually, posing a significant environmental challenge [¹]. Its derivatives are biorenewable, biodegradable, and have low toxicity, characteristics that favor its use in industrial and biomedical applications, especially when compared to synthetic materials [²-⁴].

The structure of lignin varies according to the plant species and the extraction method, generally being amorphous and rich in hydrophobic groups, which results in low solubility in various solvents [⁵]. This limitation compromises its incorporation into biomedical formulation systems. To overcome this obstacle, chemical modifications such as acetylation, esterification, nanoparticle formation, and functionalization with specific groups have been proposed [³, ⁶, ⁷]. However, such strategies, while effective in modulating physicochemical properties, tend to increase production costs, require the use of organic solvents, and may not represent sustainable or economically viable approaches for antimicrobial applications.

Lignin exhibits intrinsic biological activity, attributed to the presence of structural units such as syringyl, guaiacyl, and p-hydroxyphenyl, which contain free phenolic groups with reactive potential [⁸]. Conventional extraction methods often result in lignin with a low molecular weight and a high content of phenolic and alcoholic hydroxyl groups, which may favor its bioactivity. Still, there is room for improvement of its properties through technologies that optimize its molecular structure [⁹].

The growing antimicrobial resistance poses a significant threat to global public health, with a direct impact on morbidity, mortality, and the costs of healthcare systems [¹⁰]. Pathogens such as Candida albicans and Staphylococcus aureus have demonstrated resistance to synthetic antimicrobials, requiring the development of safer and more sustainable therapeutic alternatives [¹¹,¹²]. In this context, the search for biorenewable antimicrobial agents, aligned with the principles of green chemistry, has intensified [⁹].

Recent advances in the valorization of lignin highlight its potential as a functional material, due to its antioxidant properties, thermal stability, and antimicrobial activity [¹³]. Its use contributes to reducing dependence on synthetic inputs, mitigating environmental impacts, and valorizing industrial waste [¹⁴]. Lignin applications include controlled drug delivery systems, tissue engineering, antioxidant agents, antimicrobials, and protection against ultraviolet radiation [⁸,⁹,¹⁵-¹⁷].

Given the scarcity of studies addressing the solubilization of lignin in its natural form with a focus on antimicrobial applications, this study aimed to investigate and elucidate how different strategies for solubilizing kraft lignin influence its physicochemical properties and, consequently, its antimicrobial properties. Methodologies were employed to characterize the physicochemical properties of extracts obtained using various solubilization techniques. The analyses included determination of molecular weight, polydispersity, structural composition, phenolic hydroxyl content, and toxicity assessment. These characteristics were correlated with the antimicrobial activity profile against Candida albicans, Staphylococcus aureus, and Escherichia coli strains, given their significant clinical representativeness and impact on public health. In this context, this study contributes to the advancement of knowledge on the functional use of kraft lignin, providing support for future applications in bioactive systems and sustainable technologies.

MATERIAL AND METHODS

Materials

Kraft lignin provided by the company Suzano Papel e Celulose, absolute P.A. ethyl alcohol, tetrahydrofuran and dimethyl sulfoxide from Êxodo Científica, polystyrene (Sigma-Aldrich), sodium hydroxide (Dinâmica).

Lignin Extract Production Methods

Solubilization by magnetic stirring and temperature combined with ultrasound

One gram of raw lignin was dissolved in 100 mL of distilled water (1:100 g/mL), under heating and magnetic stirring for 1 hour. Subsequently, the solution was subjected to an Eco-Sonics ultrasonic sonicator (model QR350) for 3 minutes at a frequency of 20 kHz, using a continuous pulse with power adjusted to 385W, resulting in the LigM1 extract.

Aqueous Extraction by Decoction in a Water Bath

One hundred grams of lignin were added to 1 liter of distilled water, in a 1:10 ratio (g/mL). The lignin was placed on common filter paper with a porosity of 10 to 15 μm, sealed, and then submerged in the water. It remained in a water bath for 2 hours at an average temperature of 65°C. Finally, the filter was removed from the liquid, which was then transferred to a rotary evaporator for 2 hours at 50°C. Finally, after the aqueous lignin extract was removed from the equipment, the material was dried with hot steam to reduce the solvent, yielding a pasty material known as the LigM2 extract.

Aqueous Extraction by Decoction in an Ultrasonic Bath

One hundred grams of lignin were added to 1 liter of distilled water in a 1:10 ratio (g/mL) on a common filter paper with a porosity of 10 to 15 μm. The filter with lignin was submerged in the water, and the beaker containing the contents was placed in an Eco-sonics ultrasonic bath (model Q 1.8/40) at a frequency of 25 kHz and 66 W. The process lasted for 2 hours, with the water in the ultrasonic bath container constantly changing, aiming for an average temperature of 70°C. Subsequently, the material was subjected to vacuum filtration using a filter paper filtration system with a porosity of 0.20 μm. The combined method aimed to facilitate the interaction of ultrasonic waves with the LigM3 extract, thereby improving its solubility.

Ethanolic Extraction with Soxhlet

The methodological approach of Morais and Vieira [¹⁸] was adopted for the production of an ethanolic lignin extract, which was obtained through the Soxhlet extraction system using absolute P.A. ethyl alcohol as the solvent. Sixty grams of raw lignin were placed in the extraction cartridge, with an average porosity of 10 to 12 μm, positioned in the equipment, which had an opening at the top for the gradual addition of ethyl alcohol, until the lignin content was fully saturated. The extraction process took place over 4 hours, yielding an extract soluble in ethanol. Finally, the material was rotary-evaporated for 30 minutes and dried with hot steam, resulting in the evaporation of the solvent and the extraction in paste form, named LigM4.

Aqueous Extraction by Magnetic Stirring

To establish a comparative parameter for the efficiency of other solubilization methods, the LigM5 extract was produced. Obtained by dissolving 1g of raw lignin in 100 mL of distilled water (1:100 g/mL), the solution was homogenized under magnetic stirring for 1 hour. In this method, no heating steps were involved.

Characterization of lignin extracts

The determination of the total solids content of each extract was performed using the gravimetric evaporation and weighing method, following the results obtained by Mazar and Paleologou [¹⁹], with modifications to the method. A crucible was dried at 100°C until a fixed weight was obtained. Then, 1 mL of each extract (LigM1, LigM2, LigM3, LigM4, and LigM5) was added to a crucible, the initial content was weighed, and they were placed in an oven for 40 minutes at 70°C. Considering the pasty nature of the LigM3 and LigM4 extracts, these were dissolved in distilled water at a concentration of 20%. The percentage of total solids (TS) found in each extract was calculated from the amount of insoluble parts found in the final mass of the process, using Equation 1:

(1) $$\text{Total solids content (\%m/m)}=\text{M solids / M solution X 100}$$

The pH was measured using a benchtop pH meter (Model VZ86505AZ), and the LigM3 and LigM4 values were determined using a 20% solution. The results were expressed by identifying the mean and standard deviation of each sample.

Gel Permeation Chromatography (GPC)

Gel permeation chromatography analyses were used to compare the raw lignin with the other extracts obtained. A Shimadzu LC-10AD system was used, employing a configuration composed of three Shodex KF-806M analytical columns (10 µm, 8 mm × 300 mm) connected in series. HPLC-grade tetrahydrofuran (THF) was used as the mobile phase. Lignin samples (2 mg) were dissolved in 2 mL of HPLC-grade THF and filtered with a 0.22 μm porosity nylon filter. Five-microliter aliquots were injected into the chromatographic system, which was operated at a flow rate of 1.0 mL/min⁻at 40 °C, with detection performed using a refractive index detector (RID-10A). The calibration curve was developed using polystyrene standards (Sigma-Aldrich 76552 with molar masses ranging from 0.2 × 10³ to 10⁴ g·mol⁻¹).

Scanning Electron Microscopy (SEM)

Morphological analysis was performed on the LigM3 and LigM4 extracts using scanning electron microscopy (SEM) with the Inspect S50 instrument. A small amount of the material was added to a pre-affixed carbon tape on a stub, and the sample was viewed at 20,000x magnification.

Infrared Spectroscopy

Fourier Transform Infrared Spectroscopy (FTIR) was performed using a Shimadzu IRTracer-100 spectrometer in the wavenumber range of 400 to 4000 cm⁻¹. Approximately 5 mg of each sample was mixed with KBr and pressed.

Determination of phenolic hydroxyl groups by UV method

Following the methodology proposed by Serrano and coauthors [²⁰], a solution was prepared by dissolving 20 mg of each extract in a mixture of 10 mL of THF and 10 mL of 0.2 M aqueous NaOH. The solution was filtered through a 0.20 μm nylon filter with a porosity of 0.20 μm. A 4 mL aliquot of the initial solution was diluted in 50 mL of three different aqueous solutions: 0.2 M NaOH, pH 6 buffer, and pH 12 buffer, obtaining a final concentration of 0.08 g/L. UV measurements were performed using a Shimadzu UV-1800 spectrophotometer in the 200-800 nm range, with the lignin solution in a pH 6 buffer as a reference. The absorbance of the maxima observed at 300 nm (A300nm) and 350 nm (A350nm) was used for calculations using Equation 2.

(2) $$\left[{\mathrm{OH}}^{-}\right]=\{[0.425\times {\mathrm{A}}_{300\mathrm{nm}}]+[0.182\times {\mathrm{A}}_{350\mathrm{nm}}]\}$$

Antimicrobial Susceptibility

The antifungal and antibacterial activity of the samples was evaluated by determining the Minimum Inhibitory Concentration (MIC), the Minimum Fungicidal Concentration (MFC), and the Minimum Bactericidal Concentration (MBC).The MIC is defined as the lowest concentration of the sample capable of inhibiting 100% of visible fungal growth [²¹]. The MFC/MBC is determined as the lowest concentration that resulted in the absence of growth in the subculture after 48 hours [²²].

Determination of antifungal activity

The MIC was determined by the broth microdilution method, following the methodology described by Fontenelle and coauthors [²¹] and in accordance with the Clinical Laboratory Standards Institute protocol M27-A3 [²³]. Two Candida albicans strains were used: one was a reference strain (ATCC 90028) and the other was a clinical isolate from a blood culture (LABMIC 0102). Roswell Park Memorial Institute (RPMI) broth was used in 96-well microdilution plates, performing serial dilutions. The plates were incubated at 36°C for 24 hours. Amphotericin B, prepared in Dimethyl Sulfoxide (DMSO), was used as a positive control in the antifungal susceptibility assay. To determine the MFC, 100 µL aliquots of solution, removed from wells that did not show turbidity in the MIC assay, were subcultured in plates containing Potato Dextrose Agar medium. The plates were incubated at 28°C to observe the growth of the fungus.

ATCC standard strains, internationally recognized as a reference in microbiological testing, were used. The use of these strains ensures the reproducibility of results, allowing consistent comparisons with future studies, since they are well-characterized strains that are widely accepted in scientific research [²⁴, ²⁵].

Determination of antibacterial activity

The determination was carried out in accordance with the standards established in document M100-S, 26th edition, Performance Standards for Antimicrobial Susceptibility Testing, of the Clinical and Laboratory Standards Institute [²⁴]. The assays were performed using Gram-positive and Gram-negative bacterial strains, respectively of Staphylococcus aureus (LABMIC 6538) and Escherichia coli (ATCC 25922). Brain Heart Infusion (BHI) broth was used in 96-well microdilution plates, performing serial dilutions. The plates were incubated at 36 °C for 24 hours. To determine the MBC, 100 µL aliquots of solution, removed from wells that did not show turbidity in the MIC assay, were subcultured in plates containing BHI Agar medium. The plates were incubated at 28 °C to observe bacterial growth. Gentamicin was used as a positive control.

Cytotoxicity

The cytotoxicity of LigM2 and LigM4 was assessed using the Alamar Blue assay, as previously described [²⁶]. Initially, peripheral blood mononuclear cells (PBMCs) were isolated, washed, and resuspended in RPMI 1640 medium supplemented with 20% fetal bovine serum, 2 mM glutamine, 100 U/mL penicillin, and 100 μg/mL streptomycin at 37 °C with 5% CO₂. Phytohemagglutinin (3%) was added at the beginning of the culture. All studies were conducted in accordance with Brazilian research guidelines (Law 466/2012, National Health Council) and the Declaration of Helsinki.

Next, the cells were placed in 96-well plates at a density of 1 × 10⁶ cells/well. Samples LigM2 and LigM4 (0.75 and 1.5 mg/mL) were added to each well after overnight incubation for adhesion, and the incubation was continued for an additional 24 hours. Doxorubicin (Laboratorio IMA S.A.I.C., Buenos Aires, Argentina) at a concentration of 10 µg/mL was used as a positive control. Four hours before the end of incubation, 20 μL of resazurin (30 μM) (Sigma-Aldrich Co., St. Louis, MO, USA) was added to each well. Absorbance at 570 and 600 nm was quantified using a SpectraMax 190 microplate reader (Molecular Devices, Sunnyvale, CA, USA).

RESULTS

To elucidate the influence of solubilization methods on the properties of kraft lignin, physicochemical, structural, and biological analyses were conducted. Initially, pH and solids content were determined to evaluate acidity and solubilization efficiency. Subsequently, characterizations by GPC, SEM, FTIR, and UV-Vis allowed the investigation of molecular weight, morphology, functional groups present, and phenolic hydroxyl content. Finally, the extracts were subjected to antimicrobial assays against fungal and bacterial strains, aiming to correlate their structural characteristics with the observed biological activity. The cytoxicity of the extracts was also investigated.

Identification of pH and total solids content

The identification of the pH of the extracts obtained from the solubilization of lignin allowed the observation of the interaction between soluble and insoluble fractions, with direct implications for the bioactive properties of the samples. As shown in Table 1, all samples exhibited acidic behavior in aqueous medium.

Table 1
Total solids content and pH.

Sample	%TS	pH
LigM1	1.21%	4.14 ± 0.08
LigM2	10.39%	4.33 ± 0.3
LigM3	0.20%	5.02 ± 0.09
LigM4	12.78%	4.94 ± 0.07
LigM5	13.76%	3.98 ± 0.02

• ^*
  Total solids content (TS%) found in each mL of solution.

The LigM5 extract had a pH of 3.98, showing a greater release of H+ protons in solution. This value was the lowest among the extracts analyzed, indicating greater acidity. LigM1, obtained using a method involving ultrasonic irradiation, had a higher pH than LigM5, although it was still within the acidic range. LigM2, subjected to moderate heating, revealed an intermediate pH between the extracts. LigM3 had the highest pH (5.02), followed by LigM4 (4.94), both of which were obtained using methodologies with more intense modifications, such as decoction and ultrasonic cavitation.

Regarding the total solids content, LigM1 exhibited the lowest value at 1.21%, suggesting greater efficiency in particle dispersion. LigM2 had an intermediate content, consistent with the type of heat treatment applied. LigM3 showed the lowest percentage of insoluble fractions (0.20%), a result attributed to the action of ultrasonic cavitation, which favored solubilization. LigM4, in turn, presented a solids content of 12.78%, a high value resulting from the dilution of the pasty ethanolic extract with water to 20%, which promoted particle regrouping. LigM5 presented a concentration of 13.76%, indicating a higher concentration of insoluble material.

Gel permeation chromatography (GPC)

GPC analysis facilitated the identification and differentiation of the molecular weight and size of kraft lignins through their modifications, which were calculated by the average molecular weights (Mw) and average number (Mn), as well as the polydispersity values (PDI = Mw/Mn).

The results presented in Table 2 show that LigM2 and LigM3 exhibit similar polydispersity, with a polydispersity index of 1.08. They also have average weights of 810 and 824 g/mol, respectively, indicating a lower average molecular weight compared to the other samples. The result for LigM4 shows the highest Mw (1103 g/mol) compared to the others, suggesting that the extraction of more complex lignin fractions occurred. LigM1, with a molecular weight of 1079 g/mol, also presented a high molecular weight.

Table 2
Average values of molecular weight and polydispersity.

Sample	Mw (g/mol)	Mn (g/mol)	PDI (Mw/Mn)
LigM1	1079	934	1.15
LigM2	810	746	1.08
LigM3	824	760	1.08
LigM4	1103	935	1.18
LigM5	1021	895	1.14

• ^*
  Mw: average by weight, Mn: average by number, Mw/Mn: polydispersity.

Scanning Electron Microscopy (SEM)

SEM analysis, performed at 20,000x magnification, is shown in Figure 1, illustrating the dimensions and dispersion of LigM3 and LigM4 particles. The particle dispersion found in LigM3 suggests an increase in biological activity. In the morphology of LigM4 (Figure 1A), spherical and well-dispersed particles are shown. Apparently, a film formed when the material dried, with an irregular structure that indicated reliefs and depressions, as well as slightly visible particles of heterogeneous sizes.

Figure 1
Scanning Electron Microscopy (20,000x) of lignin (A) LigM3 and (B) LigM4.

Fourier Transform Infrared Spectroscopy (FTIR)

FTIR analysis elucidated the structural composition of the proposed materials. The spectra obtained from lignin (Lig) and extracts are shown in Figure 2. The corresponding bands were attributed to previous studies addressing the presence of syringyl (S) and guaiacyl (G) monomers in kraft lignin, as well as the presence of their functional groups [²⁷].

Figure 2
FTIR spectra of kraft lignin: LigM1(purple), LigM2(pink), LigM3(blue), LigM4(orange), LigM5(green).

All spectra show broad and intense bands between 3400-3500 cm⁻¹, observed at 3460, 3442, and 3429 cm⁻¹, characteristic of stretching vibrations of hydroxyl groups (O-H) in phenolic and aliphatic structures [²⁸]. In the aliphatic region, peaks were identified at 2940 cm⁻¹ (asymmetric stretching) and 2840 cm⁻¹ (symmetric stretching), attributed to C-H vibrations of methoxyl (-OCH₃), methyl (CH₃), and methylene (CH₂) groups [²⁹].

Prominent bands at 1610 cm⁻¹ and 1512 cm⁻¹ were attributed to C=C stretching of the aromatic ring [²⁹], while the band at 1420 cm⁻¹ is related to C-H deformations and coupled in-plane aromatic vibrations. In the fingerprint region of hardwood kraft lignin, bands were observed at 1215 cm⁻¹ (Guaiacil), 1327 cm⁻¹ and 1114 cm⁻¹ (Syringyl), in addition to a signal at 830 cm⁻¹ associated with syringyl and p-hydroxyphenylpropane units [³⁰, ³¹]. Finally, the spectrum located at 620 cm⁻¹ was observed only in LigM2, related to C-S bending, due to the use of Na₂S in the kraft pulping process [³²,³³].

Identification of the phenolic hydroxyl group by the UV method

The analysis of the UV spectra, based on the difference in absorption of the phenolic groups, shows two distinct peaks at approximately 300nm and 350nm, corresponding to the selective ionization of the phenolic groups in the aromatic system of lignin when exposed to an alkaline medium. The calculation resulting from Equation 2 shows values ranging from 1.413 to 4.475 mmol/g for the content of phenolic hydroxyls, which falls within the range reported for lignins, generally between 1-4 mmol/g [³⁴].

The data obtained, shown in Table 3, show that the content of free phenolic groups in the lignin samples varied significantly depending on the solubilization method used. Lignin extracted by direct heating (LigM1) showed the highest content, at 4.475 mmol/g, while extraction by decoction with water (LigM2) resulted in the lowest value, at 1.413 mmol/g. Ultrasound-assisted decoction (LigM3) yielded an intermediate value of 1.508 mmol/g, close to that obtained by LigM2. Soxhlet extraction with ethanol (LigM4) led to a significant increase in the content of phenolic groups, reaching 3.253 mmol/g. Finally, lignin solubilized in THF showed a value of 2.246 mmol/g, which is also an intermediate value among the other methods.

Table 3
Identification of the total content of phenolic hydroxyl groups present in each sample.

Sample	Method	A300	A350	Total OH phenolic content (mmol/g)
LigM1	Heating	1.092	0.794	4.475
LigM2	Decoction	0.289	0.381	1.413
LigM3	Ultrasound + decoction	0.275	0.485	1.508
LigM4	Soxhlet + Rotary evaporator	0.751	0.677	3.253
LigM5	Lignin+ water	0.494	0.525	2.246

Antimicrobial Sensitivity Identification

Table 4 shows the antimicrobial potential of the extracts, excluding only LigM5. The results did not indicate biological activity against the Gram-negative bacterium E. coli in any of the samples, an observation already reported previously [³⁵-³⁷]. The LigM3 extract demonstrated antifungal activity (MIC 1.25 mg/mL), which was twice as effective as LigM2 (MIC 2.5 mg/mL) for the ATCC 90028 strain. For LABMIC 0102 (MIC 1.25 mg/mL), both samples exhibited the same inhibition profile. However, neither sample showed MFC. Although LigM2 exhibited lower antifungal activity against the strain in question, it displayed an antibacterial profile against S. aureus, with an MIC of 0.625 mg/mL and an MBC of 2.5 mg/mL. In contrast, LigM3 showed no antibacterial activity.

Table 4
Antimicrobial behavior of soluble lignin fractions by the Microdilution method.

Sample	Candida albicans				Staphylococcus aureus
	ATCC 90028		LABMIC 0102		LABMIC 6538
	MIC	MFC	MIC	MFC	MIC	MBC
LigM1	N.I.	N.D.	N.I.	N.D	N.I.	N.D
LigM2	2.5 mg/mL	N.D.	1.25 mg/mL	N.D.	0.625 mg/mL	2.5 mg/mL
LigM3	1.25 mg/mL	N.D.	1.25 mg/mL	N.D.	N.I.	N.D.
LigM4	0.625 mg/mL	N.D.	0.312 mg/mL	N.D.	1.25 mg/mL	N.D.
Anph. B Gentamicin	0.5 μg/mL ------------		0.5 μg/mL ------------		------------ 0.039 μg/mL	------------- 0.156 μg/mL

LigM4 showed significantly better antifungal activity against Candida albicans strains, with MICs of 0.625 mg/mL for ATCC 90028 and 0.312 mg/mL for LABMIC 0102. For S. aureus, LigM4 had an MIC of 1.25 mg/mL, which is twice as high as that of LigM2 (0.625 mg/mL).

Cytotoxicity

The assessment of cytotoxicity in PBMCs using the Alamar Blue assay indicates that LigM2 and LigM4 showed high cell viability at the concentrations analyzed (0.75 and 1.5 mg/mL), with no evidence of toxicity. Figure 3 shows that at 1.5 mg/mL, LigM4 increased the percentage of cell viability compared to the previous concentration, while LigM2 maintained the percentage, confirming the absence of a classic dose-dependent relationship. This result may be related to the bioactivity profile due to the content of phenolic hydroxyls, known to act as antioxidants and cell modulators [⁵]. The positive control with doxorubicin (DXR) 10 µg/mL, strongly reduced cell viability, validating the sensitivity of the method [²⁶].

Figure 3
Cell viability of LigM2 (pink), LigM4 (orange) and control DXR (blue).

DISCUSSION

The data obtained reveal that solubilization methods have a significant influence on the physicochemical properties and bioactivity of kraft lignin. The acidity observed in the extracts is associated with the presence of functional groups, such as phenolics and carboxylic acids, which release protons in an aqueous medium, giving the solutions an acidic character [³⁹,⁴⁰]. The lower pH of pure lignin (LigM5) suggests a higher concentration of these groups, which may favor interactions with microbial cell membranes. According to Lourençon and coauthors [⁴¹], lignins from eucalyptus tend to precipitate in more acidic ranges, retaining oxidative groups. In addition, phenolic hydroxyls, even with slight acidity, can interfere with the integrity of the plasma membrane, promoting acidification of the medium and inhibiting H⁺-ATPase, an enzyme essential for ATP generation, thereby compromising microbial energy metabolism [³¹, ⁴²].

The increase in pH observed in LigM3 (5.02) and LigM4 (4.94) is directly associated with the more intense chemical modifications applied, such as ultrasonic cavitation and decoction. These techniques not only alter the pH, but also influence the solubility of lignin, favoring colloidal dispersion in less acidic media [⁴³]. The ultrasonic bath, for example, promotes cavitation and fragmentation of larger particles, increasing molecular mobility [⁴⁴,⁴⁵]. Pérez-Rafael and coauthors [⁴⁶] highlight that sonication intensifies dispersion and reduces particle size with increasing exposure time. On the other hand, the high solids content in LigP (13.76%) and LigM4 (12.78%) reveals limitations in solubilization. In the case of LigM4, dilution of the pasty ethanolic extract in water to 20% promoted particle regrouping, justifying the high concentration of insoluble material.

GPC analysis showed that the methods influenced the molecular weight and polydispersity of the extracts. Soxhlet extraction (LigM4) had the highest average molecular weight (1103 g/mol), suggesting the extraction of more complex lignin fractions. The extracts obtained by decoction (LigM2) and ultrasound (LigM3) had lower molecular weights, which may favor diffusion and interaction with biological targets [⁴⁷]. Although these have a lower phenolic group content, experimental studies have shown that lignin nanoparticles can more easily break through microbial membranes, relating bioactivity to their molecular structure [⁴⁷,⁴⁸].

Structural characterization by FTIR confirmed the preservation of syringyl and guaiacyl units in all extracts, indicating that the solubilization methods did not compromise the aromatic structure of lignin [³¹,⁵⁰]. UV-Vis analysis revealed significant variations in phenolic hydroxyl content, with LigM1 and LigM4 exhibiting the highest values (4.475 and 3.253 mmol/g, respectively). Active phenolic groups contribute directly to antimicrobial activity through a mechanism of action that triggers oxidative stress, damaging microbial cells, altering the permeability of the cytoplasmic membrane, and ultimately leading to the loss of essential intracellular substances, which induces apoptosis [⁵¹-⁵³].

In biological assays, the extracts were found to be selective against the microorganisms tested. No sample showed activity against Escherichia coli, and Nasrollahian and coauthors [⁵⁴] corroborate this result, stating that substances with a molecular weight greater than 600 generally cannot cross the double membrane barrier of the bacterium. In contrast, LigM4 showed the best antifungal activity against Candida albicans, with MICs of 0.625 and 0.312 mg/mL, possibly due to the high content of phenolic groups outweighing the molecular weight. LigM2 stood out for its antibacterial activity against Staphylococcus aureus, suggesting that lignin fractions with lower structural complexity and lower molecular weight may have relevant bioactive action. Although most mechanistic insights remain inferential, reviews suggest that lignin derivatives may act by rupturing the membrane and generating ROS [⁵⁵].

The cytotoxicity of the extracts, LigM2 and LigM4, proved to be relevant in PBMC at the concentrations evaluated, reinforcing the biocompatibility of the material. Phenolic compounds derived from lignin stimulate metabolic activity and protect cells against oxidative stress [³⁸,⁵⁶]. Therefore, the increase in cell viability of LigM4 at 1.5 mg/mL can be explained by the higher content of phenolic hydroxyls found when compared to LigM2. The results suggest that, in addition to being non-toxic, these extracts may exert positive bioactive effects, corroborating studies that point to lignin as a promising biomaterial for pharmacological and biomedical applications [⁵⁶].

The results presented reinforce that the choice of solubilization method is crucial for modulating the properties of lignin and its effectiveness as an antimicrobial agent. The correlation between structural characteristics and bioactivity observed in this study contributes to an understanding of kraft lignin's behavior and expands the possibilities of its use in sustainable, bioactive formulations.

CONCLUSION

The results obtained demonstrate that different solubilization methods have a direct influence on the physicochemical properties and antimicrobial activity of kraft lignin. Soxhlet extraction with ethanol yielded higher phenolic hydroxyl content, correlating with better antifungal performance against Candida albicans. On the other hand, decoction with a rotary evaporator, in addition to antifungal activity, showed superior antibacterial activity against Staphylococcus aureus. This suggests that, despite the reduced phenolic hydroxyl content, the low molecular weight facilitated the mechanism of action with the bacterial membrane. The results indicate that different solubilization approaches can direct the bioactivity of the extracts. Additionally, both methods appear to utilize non-toxic materials.

Such experimental evidence reinforces the potential of kraft lignin as a bioactive agent, especially in contexts of resistant infections that demand sustainable alternatives to synthetic antimicrobials. The detailed characterization of the extracts provides support for future investigations focused on the application of lignin in biomedical systems and microbiological control technologies.

• Funding:
  This work was supported by the Fundação Cearense de Apoio ao Desenvolvimento Científico e Tecnológico (FUNCAP) through a master fellowship awarded to Taysse H. Ferreira grant number BMD-0008-02718.01.10/24
• Institutional Review Board Statement: Not applicable.
• Informed Consent Statement: Not applicable.

Use of Generative Artificial Intelligence

The authors declare that generative artificial intelligence (AI) or AI-assisted tools were used under full human supervision. The tool(s) and version(s) used, and their purpose, are described here: The tool(s) and version(s) used, and their purpose, are described here: Consensus (web version, accessed from November 2025 to February 2026) was used for scientific literature search, and Grammarly (version current as of February 2026) was used for grammatical refinement and language editing. No confidential or sensitive data were uploaded to such tool(s), and all AI-assisted content was checked, corrected and approved by the authors, who take full responsibility for the integrity and originality of the manuscript.

Acknowledgments:

The authors are thankful for Central Analítica da Universidade Federal do Ceará (UFC) for performing the SEM (Scanning Electron Microscopy) analysis.

Data Availability Statement:

Research data are only available upon request for corresponding author.

REFERENCES

• ¹ Liu H, Dai Z, Cao Q, Shi X, Wang X, Li H, et al. Lignin/polyacrylonitrile carbon fibers: The effect of fractionation and purification on properties of derived carbon fibers. ACS Sustain Chem Eng. 2018;6(7):8554-62.
• ² Abraham B, Shakeela H, Devendra L, Arun K, Ragavan V, Brennan C, et al. Lignin nanoparticles from Ayurvedic industry spent materials: Applications in Pickering emulsions for curcumin and vitamin D3 encapsulation. Food Chem. 2024;458:140284.
• ³ Freitas F, Cerqueira M, Gonçalves C, Azinheiro S, Garrido-Maestu A, Vicente A, et al. Green synthesis of lignin nanoand micro-particles: Physicochemical characterization, bioactive properties and cytotoxicity assessment. Int J Biol Macromol. 2020;163:1798-809.
• ⁴ Alqahtani M, Alqahtani A, Kazi M, Ahmad M, Alahmari A, Alsenaidy M, et al. Wound-healing potential of curcumin loaded lignin nanoparticles. J Drug Deliv Sci Technol. 2020;60:102020.
• ⁵ Zheng L, Lu G, Pei W, Yan W, Li Y, Zhang L, et al. Understanding the relationship between the structural properties of lignin and their biological activities. Int J Biol Macromol. 2021;190:291-300.
• ⁶ Chunyan T, Haojie D, Shuo T, Liuyun J, Bingli M, Yue W, et al. A combined-modification method of carboxymethyl β-cyclodextrin and lignin for nano-hydroxyapatite to reinforce poly(lactide-co-glycolide) for bone materials. Int J Biol Macromol. 2020;160:142-52.
• ⁷ Mearaj S, Ajaz A, Kim T, Choi J. Bioactive and hemocompatible PLA/lignin bio-composites: Assessment of in vitro antioxidant activity for biomedical applications. ACS Appl Bio Mater. 2023;6(9):3648-60.
• ⁸ Lin M, Yang L, Zhang H, Xia Y, He Y, Lan W, et al. Revealing the structure-activity relationship between lignin and anti-UV radiation. Ind Crops Prod. 2021;174:114212.
• ⁹ Yun J, Wei L, Li W, Gong D, Qin H, Feng X, et al. Isolating high antimicrobial ability lignin from bamboo kraft lignin by organosolv fractionation. Front Bioeng Biotechnol. 2021;9:683796.
• ¹⁰ Belay W, Getachew M, Tegegne B, Teffera Z, Dagne A, Zeleke T, et al. Antimicrobial resistance with a focus on antibacterial, antifungal, antimalarial, and antiviral drugs resistance, its threat, global priority pathogens, prevention, and control strategies: A review. Ther Adv Infect Dis. 2025;12:20499361251340144.
• ¹¹ Gow N, Johnson C, Berman J, Coste A, Cuomo C, Perlin D, et al. The importance of antimicrobial resistance in medical mycology. Nat Commun. 2022;13:5352. Doi: https://doi.org/10.1038/s41467-022-32249-5
  » https://doi.org/10.1038/s41467-022-32249-5
• ¹² Douglas E, Wulandari S, Lovell S, Laabei M. Novel antimicrobial strategies to treat multi-drug resistant Staphylococcus aureus infections. Microb Biotechnol. 2023;16(7):1456-74. doi: https://doi.org/10.1111/1751-7915.14268
  » https://doi.org/10.1111/1751-7915.14268
• ¹³ Gordobil O, Olaizola P, Banales JM, Labidi J. Lignins from agroindustrial by-products as natural ingredients for cosmetics: Chemical structure and in vitro sunscreen and cytotoxic activities. Molecules. 2020;25(5):1131. doi: https://doi.org/10.3390/molecules25051131
  » https://doi.org/10.3390/molecules25051131
• ¹⁴ Li J, Guan D, Xia S, Fan Y, Zhao K, Zhao Z, et al. Recent advances, challenges, and opportunities in lignin valorization for value-added chemicals, biofuels, and polymeric materials. Energy Convers Manage. 2024;322:119123. doi: https://doi.org/10.1016/j.enconman.2024.119123
  » https://doi.org/10.1016/j.enconman.2024.119123
• ¹⁵ Lizundia E, Armentano I, Luzi F, Bertoglio F, Restivo E, Visai L, et al. Synergic effect of nanolignin and metal oxide nanoparticles into poly(l-lactide) bionanocomposites: Material properties, antioxidant activity, and antibacterial performance. ACS Appl Bio Mater. 2020;3(8):5263-74. doi: https://doi.org/10.1021/acsabm.0c00637
  » https://doi.org/10.1021/acsabm.0c00637
• ¹⁶ Ullah I, Chen Z, Xiea Y, Khan S, Singh S, Yu C, et al. Recent advances in biological activities of lignin and emerging biomedical applications: A short review. Int J Biol Macromol. 2022;208:819-32.
• ¹⁷ De Albuquerque T, Cavalcante V, Da Silva Rocha W, De Macedo A, Rocha M. Lignin-based hydrogels extracted from cashew bagasse and their application in antimicrobial dressings. Int J Biol Macromol. 2024;262(Pt 2):130169. Doi: 10.1016/j.ijbiomac.2024.130169
  » https://doi.org/10.1016/j.ijbiomac.2024.130169
• ¹⁸ Morais S, Vieira Í. Introduction to natural product prospecting. [S.l.: s.n.]; 2018.
• ¹⁹ Mazar A, Paleologou M. Comparison of the effects of three drying methods on lignin properties. Int J Biol Macromol. 2023;258(Pt 2):128974. Doi: https://doi.org/10.1016/j.ijbiomac.2023.128974
  » https://doi.org/10.1016/j.ijbiomac.2023.128974
• ²⁰ Serrano L, Esakkimuthu ES, Marlin N, Brochier-Salon MC, Mortha G, Bertaud F. Fast, easy, and economical quantification of lignin phenolic hydroxyl groups: Comparison with classical techniques. Energy Fuels. 2018;32(5):5969-77. doi: https://doi.org/10.1021/acs.energyfuels.8b00383
  » https://doi.org/10.1021/acs.energyfuels.8b00383
• ²¹ Fontenelle ROS, Morais SM, Brito EHS, Brilhante RSN, Cordeiro RA, Rocha MFG, et al. Chemical composition, toxicological aspects and antifungal activity of essential oil from Lippia sidoides Cham. J Antimicrob Chemother. 2007;59(5):934-40. doi: https://doi.org/10.1093/jac/dkm066
  » https://doi.org/10.1093/jac/dkm066
• ²² Fontenelle ROS, Morais SM, Brito EHS, Brilhante RSN, Cordeiro RA, Rocha MFG, et al. Antifungal activity of essential oils of Croton species from the Brazilian Caatinga biome. J Appl Microbiol. 2008;104(5):1383-90.
• ²³ Clinical and Laboratory Standards Institute (CLSI). Reference method for broth dilution antifungal susceptibility testing of yeasts; approved standard. 3rd ed. CLSI document M27-A3. Wayne, (PA): CLSI; 2008.
• ²⁴ Clinical and Laboratory Standards Institute (CLSI). Methods for dilution antimicrobial susceptibility tests for bacteria that grow aerobically. 10th ed. CLSI document M07-A11. Wayne, (PA): CLSI; 2018.
• ²⁵ American Type Culture Collection (ATCC). ATCC Standards and Reference Strains. Gaithersburg, (MD): ATCC; 2023 [cited 2026 Jan 8]. Available from: https://www.atcc.org
  » https://www.atcc.org
• ²⁶ Ahmed SA, Gogal RM, Walsh JE. A new rapid and simple non-radioactive assay to monitor and determine the proliferation of lymphocytes: an alternative to [3H]thymidine incorporation assay. J Immunol Methods. 1994;170(2):211-24. Doi: 10.1016/0022-1759(94)90396-4
  » https://doi.org/10.1016/0022-1759(94)90396-4
• ²⁷ Faix O. Classification of lignins from different botanical origins by FT-IR spectroscopy. Holzforschung. 1991;45(s1):21-7. doi: https://doi.org/10.1515/hfsg.1991.45.s1.21
  » https://doi.org/10.1515/hfsg.1991.45.s1.21
• ²⁸ Cemin A, Ferrarini F, Poletto M, Bonetto LR, Bortoluz J, Lemée L, et al. Characterization and use of a lignin sample extracted from Eucalyptus grandis sawdust for the removal of methylene blue dye. Int J Biol Macromol. 2021;170:375-89. Doi: https://doi.org/10.1016/j.ijbiomac.2020.12.155
  » https://doi.org/10.1016/j.ijbiomac.2020.12.155
• ²⁹ García-Fuentevilla L, Rubio-Valle JF, Martín-Sampedro R, Valencia C, Eugenio ME, Ibarra D. Different kraft lignin sources for electrospun nanostructures production: Influence of chemical structure and composition. Int J Biol Macromol. 2022;214:554-67. Doi: https://doi.org/10.1016/j.ijbiomac.2022.06.121
  » https://doi.org/10.1016/j.ijbiomac.2022.06.121
• ³⁰ Cherif MF, Trache D, Brosse N, Benaliouche F, Tarchoun AF. Comparison of physicochemical properties and thermal stability of organosolv and kraft lignins from hardwood and softwood biomass for potential valorization. Waste Biomass Valor. 2020;11:6541-53.
• ³¹ Eugenio ME, García-Fuentevilla L, Martín-Sampedro R, Santos JI, Wicklein B, Barra D. Tuning antioxidant and antibacterial properties of lignin by physicochemical modification during sequential acid precipitation of kraft black liquor. Wood Sci Technol. 2025;59(4).
• ³² Monteiro VAC, Silva KT, Silva LRR, Mattos ALA, Freitas RM, Mazzetto SE, et al. Selective acid precipitation of Kraft lignin: a tool for tailored biobased additives for enhancing PVA films properties for packaging applications. React Funct Polym. 2021;166:104980.
• ³³ Duval A, Lawoko M. A review on lignin-based polymeric, microand nano-structured materials. React Funct Polym. 2014;85:78-96.
• ³⁴ Goldmann WM, Ahola J, Mankinen O, Kantola AM, Komulainen S, Telkki VV, et al. Determination of phenolic hydroxyl groups in technical lignins by ionization difference ultraviolet spectrophotometry (∆ɛ-IDUS method). Period Polytech Chem Eng. 2017;61(2):93-101. Doi: https://doi.org/10.3311/PPch.9269
  » https://doi.org/10.3311/PPch.9269
• ³⁵ Maisetta G, Batoni G, Caboni P, Esin S, Rinaldi AC, Zucca P. Tannin profile, antioxidant properties and antimicrobial activity of extracts from two Mediterranean Cytinus species. BMC Complement Altern Med. 2019;19(1):82. Doi: https://doi.org/10.1186/s12906-019-2487-7
  » https://doi.org/10.1186/s12906-019-2487-7
• ³⁶ Dong X, Dong M, Lu Y, Turley A, Jin T, Wu C. Antimicrobial and antioxidant activities of lignin from residue of corn stover to ethanol production. Ind Crops Prod. 2011;34(3):1629-34. Doi:10.1016/j.indcrop.2011.06.002
  » https://doi.org/10.1016/j.indcrop.2011.06.002
• ³⁷ Nada AMA, El-Diwany AI, Elshafei AM. Infrared and antimicrobial studies on different lignins. Acta Biotechnol. 1989;9(3):295-8. Doi: https://doi.org/10.1515/9783112581841-020
  » https://doi.org/10.1515/9783112581841-020
• ³⁸ Schmetz Q, Muzyka C, Berchem T, Richel A. Monitoring technical lignin partition in aqueous/alcohol biphasic systems according to pH: Influences of the molecular structure and solvent characteristics. Green Chem. 2020;22:7031-46. Doi: https://doi.org/10.1039/d0gc01905d
  » https://doi.org/10.1039/d0gc01905d
• ³⁹ Rocha ILD, Da Costa Lopes AM, Ventura SPM, Coutinho JAP. Selective separation of vanillic acid from other lignin-derived monomers using centrifugal partition chromatography: The effect of pH. ACS Sustain Chem Eng. 2022;10(15):4913-21. Doi: https://doi.org/10.1021/acssuschemeng.1c08082
  » https://doi.org/10.1021/acssuschemeng.1c08082
• ⁴⁰ Lourençon TV, De Lima GG, Ribeiro CSP, Hansel FA, Maciel GM, Da Silva K, et al. Antioxidant, antibacterial and antitumoural activities of kraft lignin from hardwood fractionated by acid precipitation. Int J Biol Macromol. 2021;166:1532-42. Doi: https://doi.org/10.1016/j.ijbiomac.2020.11.033
  » https://doi.org/10.1016/j.ijbiomac.2020.11.033
• ⁴¹ Matos M, Claro FC, Lima TAM, Avelino F, Hansel FA, Maciel GM, et al. Acetone:Water fractionation of pyrolytic lignin improves its antioxidant and antibacterial activity. J Anal Appl Pyrol. 2021;156:105175. Doi: https://doi.org/10.1016/j.jaap.2021.105175
  » https://doi.org/10.1016/j.jaap.2021.105175
• ⁴² Pinto PIF, Magina S, Budjav E, Pinto PCR, Liebner F, Evtuguin D. Cationization of eucalyptus kraft LignoBoost lignin: Preparation, properties, and potential applications. Ind Eng Chem Res. 2022;61(10):3503-15. Doi: https://doi.org/10.1021/acs.iecr.1c04899
  » https://doi.org/10.1021/acs.iecr.1c04899
• ⁴³ Gomide RAC, De Oliveira ACS, Rodrigues DAC, Oliveira CR, Assis OGB, Dias MV, et al. Development and characterization of lignin microparticles for physical and antioxidant enhancement of biodegradable polymers. J Polym Environ. 2020;28:1326-34. Doi: https://doi.org/10.1007/s10924-020-01685-z
  » https://doi.org/10.1007/s10924-020-01685-z
• ⁴⁴ Bogdan T, Koklin A, Mashchenko N, Bobrova N, Korotaeva A, Mishanin I, et al. Experimental and simulation study of lignin depolymerization in water-ethanol suband supercritical medium. J Mol Liq. 2024;415(Pt A):126284.
• ⁴⁵ Pérez-Rafael S, Ferreres G, Kessler RW, Kessler W, Blair J, Rathee G, et al. Continuous sonochemical nanotransformation of lignin: Process design and control. Ultrason Sonochem. 2023;98:106499. Doi: https://doi.org/10.1016/j.ultsonch.2023.106499
  » https://doi.org/10.1016/j.ultsonch.2023.106499
• ⁴⁶ Su C, Wang X, Hirth K, Arvanitis M, Cao Y, Fang G, et al. Bioactive lignin from maleic acid hydrotropic fractionation: Revealing the structural-bioactivity relationship. Int J Biol Macromol. 2025;302:140519.
• ⁴⁷ Morena AG, Bassegoda A, Natan M, Jacobi G, Banin E, Tzanov T. Antibacterial properties and mechanisms of action of lignin-based nanoparticles synthesized by sonoenzymatic methods. ACS Appl Mater Interfaces. 2022;14(33):37270-9. Doi: https://doi.org/10.1021/acsami.2c05443
  » https://doi.org/10.1021/acsami.2c05443
• ⁴⁸ Kozmelj TR, Voinov MA, Grilc M, Smirnov AI, Jasiukaitytė-Grojzdek E, Lucia L, et al. Lignin structural characterization and its antioxidant potential: A comparative evaluation by EPR, UV-Vis spectroscopy, and DPPH assays. Int J Mol Sci. 2024;25(16):9044. Doi: https://doi.org/10.3390/ijms25169044
  » https://doi.org/10.3390/ijms25169044
• ⁴⁹ Huang L, Li P, Jiang K, Wu W. Investigation of the interaction mechanism between lignin structural units and enzyme. J Renew Mater. 2023;11(4):1613-26. Doi: https://doi.org/10.32604/jrm.2022.023605
  » https://doi.org/10.32604/jrm.2022.023605
• ⁵⁰ Guerra A, Filpponen I, Lucia LA, Argyropoulos DS. Comparative evaluation of three lignin isolation protocols for various wood species. J Agric Food Chem. 2006;54(26):9696-705. doi: https://doi.org/10.1021/jf062433c
  » https://doi.org/10.1021/jf062433c
• ⁵¹ García A, Erdocia X, Alriols MG, Labidi J. Effect of ultrasound treatment on the physicochemical properties of alkaline lignin. Chem Eng Process. 2012;62:150-8.
• ⁵² Kai D, Tan Y, Lee J, Loh XJ. Towards lignin-based functional materials in a sustainable world. Green Chem. 2016;18(5):1175-200. Doi: https://doi.org/10.1039/c5gc02616d
  » https://doi.org/10.1039/c5gc02616d
• ⁵³ Nasrollahian S, Graham JP, Halaji M. A review of the mechanisms that confer antibiotic resistance in pathotypes of E. coli Front Cell Infect Microbiol. 2024;14:1387497. Doi: https://doi.org/10.3389/fcimb.2024.1387497
  » https://doi.org/10.3389/fcimb.2024.1387497
• ⁵⁴ Reyes DC, Ma Z, Romero JJ. The antimicrobial properties of technical lignins and their derivatives. - A review. Polymers. 2024;16(15):2181. Doi: https://doi.org/10.3390/polym16152181
  » https://doi.org/10.3390/polym16152181
• ⁵⁵ Imlimthan S, Correia A, Figueiredo P, Lintinen K, Balasubramanian V, Airaksinen A, et al. Systematic in vitro biocompatibility studies of multimodal cellulose nanocrystal and lignin nanoparticles. J Biomed Mater Res A. 2020;108(3):770-83. Doi: https://doi.org/10.1002/jbm.a.36856
  » https://doi.org/10.1002/jbm.a.36856
• ⁵⁶ Longhin EM, El Yamani N, Rundén-Pran E, Dusinska M. The alamar blue assay in the context of safety testing of nanomaterials. Front Toxicol. 2022;4:981701. Doi: https://doi.org/10.3389/ftox.2022.981701
  » https://doi.org/10.3389/ftox.2022.981701

• Editor-in-Chief:
  Paulo Vitor Farago

• Associate Editor:
  Marcos Pileggi

Publication Dates

• Publication in this collection
  17 Aug 2026
• Date of issue
  2026

History

• Received
  18 Feb 2026
• Accepted
  11 Apr 2026