Stimuli-responsive release of covalently attached Omega-3 fatty acids from electrospun chitosan membranes

Long-chain ω-3 polyunsaturated fatty acids (ω-3 PUFAs), such as docosahexaenoic acid (DHA) and eicosapentaenoic acid (EPA), possess strong anti-inflammatory and antimicrobial properties, but their therapeutic potential is severely limited by high susceptibility to oxidative degradation. Covalent immobilization onto biopolymers using cleavable ester linkages offers a route to prevent oxidation while enabling stimuli-responsive payload delivery. In this work, a transesterification-based covalent conjugation approach was developed to graft DHA and EPA onto electrospun chitosan membranes (ESCM) to achieve controlled bioactive release tailored for wound healing. DHA and EPA were conjugated to ESCM via transesterification, and successful ester bond formation was verified using Fourier-transform infrared spectroscopy (FTIR). The degree of substitution and surface loading densities were quantified via High-Performance Liquid Chromatography (HPLC), X-ray Photoelectron Spectroscopy (XPS), and acid-base titration, while surface wettability was characterized by static water contact angle measurements. In vitro release kinetics were monitored over five days in neutral phosphate-buffered saline (PBS), acidic acetate-buffered saline (ABS), and PBS containing Pseudomonas lipase. Indirect cytocompatibility and cell migration were assessed using L929 mouse fibroblasts in transwell co-cultures evaluated with luminescence viability assays and live/dead fluorescence staining. Antimicrobial efficacy was determined against both planktonic and biofilm forms of Methicillin-resistant Staphylococcus aureus (MRSA) and Pseudomonas aeruginosa. FTIR spectra confirmed successful ester linkage of DHA and EPA onto the electrospun chitosan nanofiber surfaces. The resulting functionalized membranes displayed a hydrophobic water contact angle of 127±1°, and quantitative HPLC analysis confirmed high surface substitution with approximate loadings of ~25 µg/cm 2 DHA and ~13 µg/cm 2 EPA. Elution assays demonstrated responsive release profiles, showing no measurable lipid leaching in neutral PBS, but pronounced release under acidic conditions (ABS) and enzymatically active conditions (PBS with lipase). In cell culture assays, the conjugated membranes showed high biocompatibility by maintaining greater than 90% L929 fibroblast viability while supporting cellular migration. Furthermore, the membranes significantly suppressed planktonic growth and biofilm formation of MRSA, although no significant inhibitory activity was observed against P. aeruginosa. Covalently conjugating DHA and EPA to ESCM resolves the primary clinical limitation of PUFA oxidation while preserving their therapeutic efficacy. By leveraging localized acidic pH and bacterial lipase activity as trigger mechanisms, the scaffold facilitates on-demand, targeted release of active lipids within hostile wound microenvironments. Supported by high cytocompatibility with dermal fibroblasts and targeted bactericidal action against MRSA biofilms, this composite membrane represents a viable bioactive dressing for the management of infected and chronic wounds.

Therapeutics that are non-covalently attached to the nanofibers typically result in a burst release, which can lead to undesired high concentrations, potentially causing toxicity and limiting sustained therapeutic effects (Jannesari et al., 2011). Covalent bonds, such as ester, amide, disulfide, hydrazone, and imine, can act as cleavable linkers that facilitate controlled release in response to different stimuli (Ding and Li, 2017). Stimuli-responsive drug delivery systems can enable targeted, controlled, and on-demand delivery of drugs, enhancing therapeutic efficacy, reducing dosing frequency and side effects, and improving patient compliance (Zhang et al., 2022; Yin et al., 2013). These systems release medications in response to external or internal stimuli such as pH, enzymes, light, temperature, magnetic field, etc. (Chen et al., 2023; Jennings et al., 2025).

Chitosan has functional amine and hydroxyl groups, which could be conjugated directly or through a linker with a therapeutic compound by a stimuli-responsive amide or ester bond. For example, antibiotic levofloxacin was covalently conjugated with chitosan through a hydrolysable amide bond and electrospun with PVA to produce levofloxacin-loaded PVA/chitosan composite nanofibers for controlled slow release of antibiotic (Jalvandi et al., 2017). Antitumor drugs docetaxel and paclitaxel were conjugated with low molecular weight chitosan via a cleavable bond-amide bond for docetaxel and an ester bond through succinate linker for paclitaxel, for oral delivery. Conjugated compounds enabled the prolonged, slow release of the drugs in simulated intestinal fluid (Lee et al., 2009; Lee et al., 2008). Another study reported linking ibuprofen with chitosan through an amide bond in polyethylene-grafted-chitosan and PLGA composite nanofiber, which reduced the burst release of the drug (Ji et al., 2004).

Omega-3 fatty acids are one type of polyunsaturated fatty acids (fatty acids containing more than 2 double bonds) that are essential for various body functions, particularly for brain functions, and are associated with other health benefits including antiarrhythmic, antithrombotic, and anti-inflammatory (Covington, 2004). Omega-3 polyunsaturated fatty acids (ω-3 PUFA) are named based on the position of the first double bond counting from the terminal methyl group (Figure 1). They are not synthesized within the body and must be consumed through food or supplements (Covington, 2004). ω-3 PUFA alone or in combination with other drugs have been used in treating hypertension (Cicero et al., 2009; Bercea et al., 2021), hyperlipidemia (Micallef and Garg, 2009; Backes et al., 2016), and other inflammatory diseases like psoriasis (Rahman et al., 2013), rheumatic arthritis, (Akbar et al., 2017), inflammatory bowel disease (Barbalho et al., 2016). A combination of arginine and ω-3 PUFA supplements has been found to have positive effects on wound healing rates and resistance to infection (Alexander and Supp, 2014). ω-3 PUFA oil-loaded alginate/chitosan nanocapsule was incorporated with cellulose wound bandages to improve healing (Sedik et al., 2022).

ω-3 PUFA, especially long chain ones, are susceptible to oxidation when exposed to light, environmental oxygen or high temperature limiting their use in biological applications (Ismail et al., 2016). The primary oxidation products are lipid peroxides, which later convert into byproducts like radicals, leading to loss of their activity and an off-form flavor (L et al., 2011; Miyashita, 2014). To maximize effects, ω-3 PUFA are typically encapsulated into emulsions, liposomes, microcapsules, and liquid crystalline to prevent their oxidation (Homroy et al., 2024; Du et al., 2022). These encapsulation systems are generally stabilized by adding antioxidants. Hamed et al. developed an edible curcumin loaded chitosan/alginate microsphere to encapsulate ω-3 PUFA. Curcumin was used as an antioxidant to protect the ω-3 PUFA from oxidation (Hamed et al., 2020). DHA loaded PLGA/chitosan nanoparticles can improve the stability of DHA and prevent its oxidation or degradation in the gastrointestinal tract. In addition, the nanoparticle also has the potential of slow-release effect to enhance the digestion and absorption of DHA (Liu et al., 2021). β-Lactoglobulin/oleic acid modified chitosan oligosaccharide nanoparticles have been used to entrap DHA to prevent it from oxidation (Ha et al., 2018).

Electrospun chitosan membranes act as good wound healing templates with excellent breathability, and moisture absorption capability, as well as due to their spatial structure which mimics the extracellular matrix which facilitates cell attachment and proliferation (Kamble et al., 2017; Yu et al., 2009; Son et al., 2014). ω-3 PUFA can be covalently linked to chitosan nanofibers through a cleavable ester bond for controlled release and prevention of oxidation, while retaining chitosan’s free amine group to preserve its inherent properties. Ester bond hydrolyses in response to pH changes or hydrolytic enzyme such as esterase or human serum albumin (Bender and Turnquest, 1957; Stefanidis and Jencks, 1993; Martin, 1967). The pH of normal human skin lies between 4.5–5.5 (Lukić et al., 2021). Ester bonds will allow controlled release of the acids in biologically low pH environments or infected environments in the presence of the bacterial lipase enzyme.

We developed a covalent conjugation technique of ω-3 PUFA, mainly DHA and EPA, with electrospun chitosan membrane through a transesterification reaction. In this study, we evaluated the covalent conjugation technique based on degree of substitution of EPA and DHA, the release kinetics of fatty acids in response to acidic pH and pseudomonas lipase, cytocompatibility, and antimicrobial activity. ω-3 PUFA conjugated membranes can be used as a wound dressing for sustained delivery of ω-3 fatty acids in active form to promote healing.

Materials

Chitosan (86% DDA, MW = 664.7 kDa, intrinsic viscosity = 6.249 dL/g) was purchased from ChitoLytic Inc (ON, Canada). Trifluoracetic acid (TFA), dichloromethane (DCM), triethylamine (TEA), Amano lipase from Pseudomonas fluorescens were purchased from Sigma Aldrich (MO, United States). Pyridine and acetone were purchased from Fisher Scientific Inc (NH, United States). Nordic Naturals ω-3 triglycerides liquid was purchased from Amazon, which contained 32% Docosahexaenoic acid, 48% eicosapentaenoic acid, and 20% other ω-3 PUFA.

Conjugation of ω-3 PUFA to electrospun chitosan membrane (ESCM)

Electrospun chitosan membranes (ESCMs) were fabricated using a 5.5 wt% solution of 86% degree of deacetylation (DDA) chitosan dissolved in a 70:30 (v/v) mixture of trifluoroacetic acid (TFA) and dichloromethane (DCM). The solution was loaded into a 10 mL syringe equipped with a 20-gauge, 3.81 cm blunt needle and mounted on a syringe pump. Electrospinning was performed at room temperature inside a ventilated Plexiglas enclosure maintained at 40%–60% relative humidity.

The solution was dispensed at a flow rate of 15 μL/min under an applied voltage of 26 kV (Gamma High Voltage Research, FL). Randomly oriented fibers were collected on an aluminum foil-covered, 15 cm diameter plate rotating at 8.4 rpm, positioned at a tip-to-collector distance of 15 cm. To achieve a target membrane thickness of approximately 0.7 ± 0.1 mm (700 µm), three consecutive 10 mL volumes of the chitosan solution were spun. Following fabrication, the membranes were sectioned into 10 mm diameter discs for subsequent experiments.

To remove residual TFA anions, the as-spun ESCM discs were immersed in a 10% (v/v) triethylamine (TEA) in acetone solution for 24 h under mild stirring, followed by three consecutive acetone washes. The neutralized membranes were then functionalized with ω-3 polyunsaturated fatty acid (PUFA) triglycerides. Pyridine was added to the membranes at a ratio of 10 g membrane/L. Omega-3 oil was then added slowly to the mixture at a 1:4 (v/v) ratio of ω −3 oil to pyridine under continuous stirring. The reaction was allowed to proceed overnight (24 h) at 23 °C (Figure 2). Following functionalization, the membrane discs were removed, washed with acetone (2.5 g membrane/L) for 10 min, and subsequently washed three times with deionized water at 24-h intervals. The modified ESCMs were dried in a biosafety cabinet. After membranes were fabricated, they were sectioned into 10 mm diameter discs for use in experiments.

The reaction was confirmed by Fourier Transform Infrared Spectroscopy (FTIR). FTIR spectra was recorded by attenuated total reflection technique (PerkinElmer Frontier Universal ATR, MA, United States) with a diamond crystal in the range of 500 cm -1 to 4,000 cm -1 and at a resolution of 4 cm -1. Water contact angle PUFA-conjugated was determined using VCA Optima equipment (AST products, INC, United States).

Scanning electron microscopy (SEM) images of the ESCMs were acquired using a Nova NanoSEM 650 (FEI™, Hillsboro, OR, United States). To enhance visual identification of bacterial morphology, microorganisms were identified by characteristic shape and size, and subsequently false-colored using Adobe Photoshop (San Jose, CA).

Total loading calculation

The total ω −3 fatty acids conjugated to the membrane were calculated by measuring the concentrations of DHA and EPA in the deacylated solution of the ω-3-conjugated membrane. Membranes were deacylated following the method described in Wu et al. (Wu et al., 2014). Briefly, 10 mg of ω-3 membrane was placed in a flask containing 5 ml of 5% KOH solution and stirred at 60 °C for 1 h ω-3 fatty acids was extracted from the KOH solution into an organic layer using Bligh and Dyer method (Breil et al., 2017). A 1:2 volume of methanol and chloroform solution was added to the ω-3 fatty acid-KOH solution and mixed well. Then an equal volume of water was added into the mixture, vortex and centrifuged. Fatty acid content was extracted in the lower chloroform layer, which was separated from the top layer, allowed to dry by evaporating the chloroform solvent. Later, a certain volume of ethanol was added to the dry fatty acids and concentration was determined using high performance liquid chromatography (HPLC, ThermoScientific Dionex Ultimate 3,000 Series). The column was a BDS Hypersil reversed-phase C18 column (particle size of 5 µm) (250 × 4.6 mm) (run time = 6 min, injection volume = 10 μL, wavelength = 215 nm). The mobile phase consisted of 72% acetonitrile and 0.1% orthophosphoric acid. DHA and EPA content in the reacted ESCM was determined from the HPLC reading.

Degree of substitution

The degree of substitution of DHA and EPA in the ω-3- conjugated membrane was estimated using a titration method (Wu et al., 2014; Elomaa et al., 2004). Approximately 10 ml of 75% ethanol solution was added to 340 mg of ω-3 membrane and heated in a water bath at 50 °C for 30 min. Ten ml of 0.5 N KOH solution was added to the mixture, following cooling down to room temperature, and stirred for 72 h. All the conjugated ω-3 fatty acids are released from the membrane through complete basic hydrolysis of the ester linkages and are neutralized by KOH. Any excess KOH in the mixture was estimated by titrating with a 0.5 N HCl solution. A blank solution of 75% ethanol was processed in the same way to define the control. The total mol of conjugated ω-3 fatty acids is calculated from Equations 1–3.

Where, n D H A a n d n E P A are the moles of DHA and EPA released from the ω-3 membrane, respectively, and the ratio between them was estimated from the HPLC data. x and y are the degree of substitution of DHA and EPA respectively, m D H A, m E P A and m P U F A _ E S C M are weight of DHA, EPA content and ω-3 conjugated membrane. M W D H A, M W E P A, M W c h i t o s a n are molecular weight of DHA, EPA and an average monomer unit of DDA 86% chitosan.

Surface fraction (Φ s)

X-Ray Photoelectron Spectroscopy (XPS) was used to provide an estimate of the elemental composition of the ω-3 membrane from the top layer, approximately 5 nm thick.

The Surface fraction (Φ s) of the DHA and EPA (molecular percentage of the surface occupied by fatty acid chains versus unmodified glucosamine units) was estimated from elemental composition using Equation 4:

Where, C and N are the atomic percentages of carbon and nitrogen respectively found from XPS, x and y are the degree of substitution of DHA and EPA respectively, and p and q are the number of carbon atoms in DHA and EPA. The ratio of DHA and EPA degree substitution (x/y) was estimated from HPLC loading data.

Elution

A 5-day elution profile of the EPA and DHA was evaluated in ABS, pH = 4.5 and PBS, pH = 7.4, with and without the presence of the 2 mg/ml pseudomonas lipase enzyme. To determine the release profile, ESCMs (1 cm discs) were placed in 1 ml of PBS, ABS, and PBS containing lipase (n = 4) for each condition, and the eluates were collected and replaced with fresh saline every day. The eluted ω-3 fatty acids were extracted from the buffer saline using Bligh and Dyer method. Concentrations of EPA and DHA in the extracted samples were measured by high-performance liquid chromatography.

Cytocompatibility

An indirect cytocompatibility of ω-3 PUFA conjugated ESCM release and its material was evaluated using mouse fibroblast cell lines (L929). Cells were seeded at a concentration of 10 4 cells/cm -2 in a 24-well plate in Gibco Minimum Essential Medium α (MEM α), high glucose supplemented with 10% fetal bovine serum (FBS, Gibco), and 2% (100 μg mL −1) Normocin (InvivoGen, San Diego, CA, United States) and cultured until they were confluent. Membranes were placed into trans wells, and plates were incubated at 37 °C with 5% carbon dioxide (CO 2). After 24 h, the membranes were removed and Cell viability was determined using CellTiter-Glo® (Promega, Madison, WI, United States) and expressed as a percentage of no-treatment control. The cells were stained with ThermoFisher Live/Dead Viability/Cytotoxicity kit to highlight the live and dead cells in wells treated with each group.

Antimicrobial efficacy

The antimicrobial efficacy of ω-3 PUFA conjugated ESCM was compared to traditional gauze dressing against the two most prevalent wound pathogens- Methicillin-resistant Staphylococcus aureus (ATCC 33591) and Pseudomonas aeruginosa (PAO1, wound-isolated strain) (Y et al., 2014). Membranes and gauze were incubated with approximately 1 × 10 5 colony-forming units of bacteria at 37 °C for 72 h. Following the incubation, the treatments were removed and washed three times with phosphate buffer saline before sonication to detach the bacteria from the membrane and gauze into the sterile tryptic soy broth (TSB). Biofilm growth on the membrane and gauze was then quantified using the bacteria in the TSB solution. The bacterial viability in the planktonic and biofilm form was estimated using BacTiter-Glo TM microbial cell viability assay.

Statistical analysis

Statistical analysis was performed using GraphPad Prism 10.4.2 software. The L929 cell viability in presence of ω-3 PUFA conjugated ESCM was compared to no-treatment control using a t-test (n = 5, α = 0.05). Planktonic growth (n = 5), biofilm growth on membrane (n = 4) and culture plate (n = 5) of MRSA and PAO1 were assessed first by performing the Shapiro-Wilk normality test, followed by the Brown-Forsythe equal variance test. If both passed, data was further analyzed with a one-way analysis of variance (ANOVA) followed by Holm-Sidak’s post hoc analysis to detect significance between experimental groups (α = 0.05). If normality and equal variance were not passed, data was analyzed using Kruskal–Wallis ANOVA on ranks, followed by Tukey post hoc test.

Covalent conjugation of ω-3 PUFA with ESCM

FTIR spectra reveal two newly formed peaks at 1740 cm -1 corresponding to the ester C=O stretching (highlighted in blue), and a broad peak around ∼2,900 cm -1 attributed to the alkyl group of the conjugated omega-3 (highlighted in green) in the ω-3 PUFA conjugated ESCM compared to the unreacted ESCM (Figure 3). Water contact angle of the PUFA conjugated membrane was 127° ± 1°.

Membranes contained approximately 25 μg/cm 2 of DHA and 10 μg/cm 2 of EPA, as found from the HPLC reading of deacylated ω-3 membrane. The degree of substitution of DHA and EPA based on the surface of the membrane and the total membrane is presented in Table 1. The degree of substitution of DHA and EPA on the surface of the membrane, per chitosan molecule was 1.23 and 0.71 respectively. Considering a maximum degree of substitution (DS) of 2 in the two-hydroxyl groups of chitosan, this corresponds to 61.5% DHA and 35.5% EPA substitution, resulting in a total ω-3 DS of 1.97, or an overall substitution degree of 97%. Based on the entire membrane, the degree of substitution of DHA and EPA per chitosan molecule was 0.085 (4.25%) and 0.05 (2.5%), respectively. So, the total degree of acylation based on the surface and the total material was 1.97 (97%) and 0.135 (6.75%), respectively.

Esterification allows slow and stimuli-responsive release of PUFA

HPLC data indicate pH- and lipase-sensitive release of DHA and EPA from the ω-3 PUFA-conjugated membrane (Figure 4). No release of ω-3 fatty acids was observed in neutral pH PBS, whereas significant release occurred in acidic acetate buffered saline (ABS) and in PBS containing Pseudomonas lipase. EPA release was minimal, with only a few micrograms detected on days one, two, and four, totaling approximately 20% of the initially loaded amount. In contrast, DHA showed a more consistent release profile, with an average of 20 µg released per day in both ABS and lipase-containing PBS. The PUFA membranes sustained release over 5 days in ABS, with complete DHA release achieved by day four. Overall, DHA exhibited a more stable and prolonged release, particularly in ABS.

The morphology of the PUFA-conjugated membrane was examined using SEM both before and after 5 days of elution (Figure 5). The membranes after elution were immersed in ABS and PBS containing lipase for approximately 5 weeks.

L929 cells on ω-3 PUFA conjugated membranes exhibit 90% viability

The L929 cells exhibited a viability of over 90% in indirect contact of the ω-3 PUFA-conjugated membrane using transwell, with the majority appearing green—indicating live cells—as illustrated in Figure 6.

ω-3 PUFA-conjugated membrane possesses antimicrobial activity against gram-positive and gram-negative bacterial pathogens

The ω-3 PUFA-conjugated membrane inhibited planktonic MRSA growth by approximately 75% and inhibited biofilm formation on the culture plate by around 50%, compared to the untreated control (Figure 7). Additionally, it significantly suppressed biofilm growth on the membrane surface itself, showing nearly 50% inhibition relative to the gauze control. These preliminary results highlight the effectiveness of ω-3 conjugated membrane against MRSA. While its activity against P. aeruginosa was not as notable as against MRSA, it still outperformed gauze, inhibiting nearly 40% of planktonic PAO1 and around 15% of biofilm formation on the culture plate—compared to just 8% and 0% inhibition by gauze, respectively. Moreover, the membrane inhibited approximately 25% of biofilm growth on its surface compared to the gauze surface.

SEM images of the MRSA and PAO1 attached on the gauze surface and PUFA membrane were shown in Figure 8. These images illustrate that gauze has significantly more MRSA adhered to it compared to PUFA membrane, while both gauze and PUFA membrane have similar level of PAO1 attachment.

ω-3 fatty acids, especially DHA and EPA, are explored in various formulations to promote wound healing through different mechanisms, including modulation of inflammation, promotion of early epithelialization, and reduction of excessive collagen deposition (Covington, 2004; Alexander and Supp, 2014; Severing et al., 2022). Due to their high susceptibility to oxidative degradation, ω-3 fatty acids are typically used in more stable forms such as monoglyceride, ethyl ester, or triglycerides (Shahidi and Zhong, 2010; Chevalier et al., 2021). In this study, we explored a covalent conjugation method to bind ω-3 fatty acids with chitosan nanofibers, aiming to create an effective wound healing scaffold capable of controlled, on-demand delivery of bioactive ω-3s under infected environments. This on-demand, controlled release system can prevent wound infections over an extended period, reducing the need for frequent dressing changes—which disrupt the healing process and may be inconvenient in resource-limited or austere environments.

SEM analysis revealed that the nanofibrous architecture remained relatively stable in PBS with lipase, whereas significant structural breakdown occurred in the acidic acetate buffer. This rapid degradation in acidic conditions is likely driven by the synergistic effect of chitosan’s inherent solubility at low pH and the localized release of ω-3 fatty acids. While the loss of fiber morphology is evident in acidic environments, the timing of this degradation is critical to its clinical utility. For wound dressings, this breakdown occurs following the primary therapeutic release of the ω-3 PUFAs, aligning with the end of the dressing’s functional lifespan (He et al., 2026; Dhivya et al., 2015). For regenerative scaffolds, the gradual loss of structure allows for a controlled transition where the scaffold provides essential mechanical support during the initial stages of tissue repair, before being replaced by the host’s developing extracellular matrix (Tajvar et al., 2023; Santhamoorthy and Kim, 2025; Zhang et al., 2014). Consequently, the observed degradation profile supports the material maintaining its integrity during the active treatment phase and clearing or being removed from the site once its biological function is complete. A primary limitation of this study, however, is that degradation was evaluated purely via qualitative electron microscopy rather than quantitative mass-loss tracking over time. Furthermore, these evaluations were restricted to an isolated in vitro stability model. In a physiological environment, the complex milieu of endogenous enzymes, cellular activity, and the dynamic microenvironment of wound fluid may significantly accelerate or decelerate matrix clearance compared to these controlled laboratory conditions. Future preclinical in vivo studies are required to establish definitive degradation profiles and confirm how these morphological changes correlate with active tissue healing. FTIR analysis confirmed the successful conjugation of ω-3 fatty acids onto the ESCM via ester bonds. However, quantitative assessments revealed a stark divergence between surface-specific and bulk characterization metrics. XPS analysis indicated an exceptionally high apparent surface substitution (Φs) of 1.94, whereas the back-titration method yielded a much lower bulk stoichiometry (DS bulk) of approximately 0.1. This discrepancy is a direct consequence of the core-shell functionalization pattern and the differing probing depths of the two techniques. While back-titration measures the average chemical composition across the entire volume of the 100 nm diameter fibers, XPS is strictly surface-sensitive, probing only the outermost 3–5 nm (Freire et al., 2006). Steric and geometric constraints further explain this interfacial concentration. Polyunsaturated fatty acids (PUFAs) like DHA and EPA are highly bulky, with DHA occupying a molecular volume of approximately 355 Å 3. Because the crystalline spacing of aligned chitosan monomeric units is roughly 5.2 Å, covalent conjugation forces these massive fatty acid chains to extend outward, creating a dense hydrophobic canopy (spanning roughly 1.3–2.6 nm in thickness) at the fiber interface. Because this canopy dominates the narrow 3–5 nm sampling depth of the XPS photoelectrons, the carbon-to-nitrogen ratio is heavily skewed toward the modifier, yielding a high apparent surface value. Consequently, the titration result of DS ∼ 0.1 provides a more realistic representation of the overall fiber composition. This value corresponds to approximately 10% of the total chitosan monomeric units being successfully conjugated with a PUFA molecule, translating to roughly 20% of the overall membrane weight. It is worth noting, however, that the titration value itself may represent an overestimation; if residual acetic acid remained in the matrix, it would consume a portion of the alkaline titrant, thereby artificially inflating the calculated bulk DS.

The release profiles differed markedly between the two fatty acids. EPA release was highly limited and sporadic, occurring only on days 1, 2, and 4, with a minimal yield of approximately 1 μg at each time point. Conversely, DHA exhibited a continuous, sustained release profile over a 5-day period. Intriguingly, the cumulative amount of DHA recovered during the release study exceeded its theoretically calculated initial loading, occasionally surpassing 100% of the expected yield. Rather than a statistical anomaly, this discrepancy is highly indicative of an underestimation of the baseline DHA loading. Because ω-3 fatty acids are highly susceptible to thermal degradation, a significant fraction of the DHA may have degraded during the high-temperature deacylation process required for baseline quantification, masking the true extent of the initial conjugation (Nagy et al., 2024). Ultimately, the successful release of PUFAs under acidic conditions and in the presence of lipase underscores the potential of these functionalized membranes for targeted, responsive biomedical applications. Intact, healthy skin maintains a slightly acidic environment (pH ∼ 4.7), whereas infected or chronic wounds typically shift toward alkalinity and exhibit elevated levels of bacterial lipases (Das and Bal, 2024; Chandra et al., 2020; Jaeger et al., 1994). Because these extracellular enzymes specifically hydrolyze ester linkages, the PUFA-conjugated ESCM holds strong promise as an on-demand, smart drug delivery system capable of releasing therapeutic ω-3 fatty acids directly upon contact with an active wound infection (Singh et al., 2019).

According to ISO 10993–5, a cell viability of 70% or higher in direct contact or indirect contact through agar or filter diffusion, or in presence of material extract is considered indicative of cytocompatibility (Thangaraju and Varthya, 2022). Since the mouse fibroblast cells used in this study exhibited over 90% viability when in indirect contact with PUFA conjugated membranes, the membranes were judged to be cytocompatible. ω-3 polyunsaturated fatty acids (PUFAs) are well-established modulators of inflammatory cell signaling (Tsamesidis and Kalogianni, 2023) and have demonstrated clinical efficacy in reducing cutaneous inflammatory responses and systemic oxidative stress (Lin et al., 2021). In biomaterial applications, structurally similar electrospun nanofibers incorporating ω-3 fatty acids have been shown to support the sustained growth of neuroblastoma and glial cells over a 72-h culture period (Arancı et al., 2026). Furthermore, nanoliposomal vehicles loaded with ω-3 fatty acids exhibit excellent cytocompatibility with macrophages while actively promoting cellular lipid uptake (Shabana et al., 2024). When applied directly to primary dermal lineages, such as keratinocytes and fibroblasts, unencapsulated ω-3 fatty acids elevate baseline metabolic activity, though they do not directly accelerate proliferation or migratory kinetics (Severing et al., 2022). The slow and controlled release of ω-3s from the membranes in the present study would be supportive of dermal applications, since findings from gelatin and carboxymethyl cellulose hydrogels showed lower loading concentrations of ω-3 fatty acids supported fibroblast viability, whereas elevated concentrations induced distinct cytotoxicity and hemolysis (Zamani et al., 2024). A primary limitation of the current evaluation is the reliance on a single fibroblast cell line paired exclusively with endpoint viability assays. Comprehensive in vitro profiles utilizing multi-lineage cultures and subsequent preclinical in vivo wound models are required to definitively characterize the mechanical and biological impacts of these functionalized membranes on the coordinated cellular cascade of tissue repair.

ω-3 fatty acids are well known for promoting inflammatory cell signaling and have been shown in human trials to support reduced skin inflammatory response and oxidative stress. Similar electrospun fibers with ω3 incorporated nanofibers were shown to support growth of neuroblastoma and glial cell growth over a 72-h period. Other studies showed that nanoliposomes loaded with ω-3 fatty acids were compatible with macrophages and promoted uptake of fatty acids into macrophages. For dermal cells like keratinocytes and fibroblasts, direct addition of ω-3 fatty acids increased metabolic activity but did not directly stimulate proliferation or migration. Gelatin and carboxymethyl cellulose gels loaded with varying concentrations of ω-3 fatty acids showed support of fibroblast viability at lower loading concentrations, but cytotoxicity and hemolysis at high ω-3 loading concentrations. A limitation of this study is that we used a single fibroblast cell line and only viability determination. Future in vitro work and preclinical wound models are needed to determine the effects of our membranes on cells involved in the wound healing process.

PUFA membranes had antimicrobial activity against a common wound pathogen, MRSA. Ωinhibiting more than 50% of MRSA, both in planktonic and biofilm form. Given the absence of antibiotics and high bacterial load of 1 × 10 5 colony-forming units, this level of inhibition is noteworthy, especially considering that fewer bacteria would typically be present in clean or contaminated wounds (Bessa et al., 2015). In one study, DHA and other ω-3 fatty acids showed antibacterial and anti-biofilm effects against Gram-positive E. faecium (Wei et al., 2023). DHA may also increase the susceptibility of bacteria to other antimicrobials as demonstrated in a study where DHA-loaded chitosan/alginate nanoparticles improved the biocidal activity of amoxicillin against Helicobacter pylori in an in vivo rat model (Khoshnood et al., 2023). The study suggested that the antibacterial activity of long-chain ω-3s was attributed to their ability to disrupt bacterial cell membranes and stimulate macrophage-dependent clearance, immunomodulatory, and anti-inflammatory effects. However, in this study PUFA-conjugated membranes were less active against P. aeruginosa, inhibiting around 15%, with no statistically significant differences from the control. This lower activity toward Gram-negative strains was also observed in previous studies and attributed the outer membrane, which restricts the diffusion of hydrophobic compounds through its lipopolysaccharide layer (Coraça-Huber et al., 2021; Clare et al., 2024). There is some evidence that while ω-3 fatty acids may not have strong antimicrobial activity against Gram-negative microorganisms like P. aeruginosa, they may reduce toxin formation and assist with immune cell clearance of infections (Das, 2018; Wu et al., 2016) Non-covalently attached DHA delivered from chitosan/alginate vehicle was found to have antibiofilm activity against P. aeruginosa both in vitro and in vivo, to reduce its pyocyanin production, and to accelerate burn wound healing in an in vivo rat model (Ghaneialvar et al., 2023). While these findings demonstrate the potential of polyunsaturated fatty acid (PUFA)-conjugated membranes, several limitations remain to be addressed in future work. First, the biochemical mechanisms underlying the inhibition of both planktonic and biofilm-associated microorganisms were not fully explored. Second, the current evaluation was confined to an in vitro model utilizing only two representative microbial strains, which may not capture the complex, multi-species environments typical of in vivo settings. Finally, this protocol did not evaluate the performance of the membranes in combination with additional antimicrobials routinely co-administered during clinical wound care. Addressing these factors in subsequent studies will be essential to comprehensively establish the translation and utility of PUFA-modified matrices for wound healing and alternative biomedical applications.

In summary, this study successfully demonstrated a covalent conjugation strategy to anchor ω-3 polyunsaturated fatty acids (PUFAs)—specifically DHA and EPA—to electrospun chitosan membranes, establishing a versatile, stimuli-responsive platform for localized, antibiotic-free wound therapeutics. Structural characterization via FTIR, XPS, and back-titration confirmed successful esterification and revealed a distinct core-shell functionalization architecture. While bulk loading accounted for roughly 20% of the total membrane weight, steric constraints forced the bulky PUFA chains to organize into a dense, hydrophobic canopy at the outermost 3–5 nm interface. The functionalized matrices exhibited excellent application-specific degradation profiles, maintaining stability in standard physiological conditions but undergoing rapid morphological breakdown under acidic and lipase-rich environments that mimic active wound infections. Notably, DHA demonstrated a highly favorable, continuous, and sustained 5-day release cascade. Furthermore, the membranes proved highly cytocompatible, supporting over 90% mouse fibroblast cell viability in indirect contact assays. The controlled release of localized ω-3 fatty acids provides a safe metabolic baseline for dermal lineages without risking the cytotoxicity or hemolysis associated with burst-release hydrogels. Therapeutically, the membranes demonstrated robust utility by inhibiting over 50% of the major wound pathogen MRSA in both its planktonic and biofilm phenotypes without driving antibiotic resistance.

However, several limitations remain to be addressed in future work. While the matrix was highly active against Gram-positive MRSA, the hydrophobic lipopolysaccharide outer membrane of the Gram-negative P. aeruginosa limited direct inhibitory efficacy. Additionally, structural degradation kinetics remain unquantified by formal mass-loss tracking over time, and baseline loading parameters for the highly heat-sensitive ω-3 chains were likely underestimated due to thermal degradation during the high-temperature deacylation assays required for baseline quantification. Finally, the biological evaluations were restricted to an isolated in vitro stability model utilizing a single fibroblast lineage and only two representative microbial strains, which do not fully capture the complex, multi-species biofilms, dynamic enzymatic milieus, or cellular cascades characteristic of in vivo clinical environments. Future investigations will focus on exploring the precise molecular mechanisms of microbial inhibition, evaluating performance in combination with co-administered clinical antimicrobials, and validating matrix degradation and accelerated tissue repair within preclinical in vivo wound healing models.