A Multifunctional Bioactive Nanoscale Coating Deposited by Atmospheric Pressure Plasma Polymerization of Peppermint Essential Oil

ABSTRACT

Implanted and indwelling medical devices remain challenged by infection, oxidative stress, and chronic inflammation, underscoring the need for multifunctional surface coatings to holistically address these complications. Peppermint essential oil is inherently antibacterial, antioxidant, and anti‐inflammatory, yet its integration into stable, contact‐active coatings is limited by fabrication constraints. Here, we present a one‐step atmospheric pressure plasma polymerisation process that converts peppermint essential oil into a conformal, cross‐linked coating that preserves precursor‐derived functional groups that drive broad bioactivity. While the coating is substrate‐independent, we evaluate its bioactive performance within the context of bladder catheterisation as a pilot application. It scavenges up to 90% of reactive species, reduces pro‐inflammatory cytokine expression by up to 60%, and increases anti‐inflammatory cytokines by up to 50%, while promoting macrophage polarisation toward an M2 phenotype. The coating exhibits intrinsic antibacterial activity, reducing viable bacteria by 90% (Live/Dead) and 70% (CFUs), attributed to membrane disruption of Gram‐negative pathogens. In turn, this interaction potentiates the activity of colistin and levofloxacin, two antibiotics used in catheter‐associated urinary tract infection management. Together, these findings establish a stable, multifunctional coating capable of mitigating infection, alleviating inflammation, and enhancing antibiotic performance, while offering a sustainable route for essential‐oil‐derived biomaterials.

Atmospheric plasma polymerization transforms peppermint essential oil into ultrathin, substrate‐independent bioactive coatings with antioxidant, contact‐active antibacterial, and immunomodulatory functions. The coating disrupts Gram‐negative pathogens, potentiates prophylactic antibiotics, suppresses pro‐inflammatory signaling, and promotes M2 macrophage polarization. These sustainable, multifunctional films offer a promising strategy for improving outcomes of implantable medical devices, including urinary catheters.

1. Introduction

Implantable and indwelling medical devices are indispensable in modern healthcare. However, the process of introducing foreign materials into the host physiological environment can trigger complications, including infection [1, 2], inflammation, pain, impaired tissue healing, and in severe cases, sepsis and death [3]. Even in the absence of infection, mechanical forces associated with device placement can trigger a cascade of inflammatory responses [4], including the recruitment of effector cells such as neutrophils and macrophages [5], which generate reactive oxygen species (ROS) as part of their defence function [6]. Excessive ROS leads to oxidative stress, exacerbating tissue damage, delaying healing, and impairing device integration [7]. To address these complications, a range of surface modification approaches has been developed, aiming to improve healing [8, 9], reducing oxidative stress [10], alleviating inflammation [11], or inhibiting bacterial colonization [12, 13]. While individually effective, an ideal biomaterial coating should combine all these properties into a single, multifunctional coating to address a diverse range of complications holistically, thereby simultaneously resolving inflammation and preventing infection.

Plant‐derived essential oils are attractive to fill this purpose because they contain a diverse composition of bioactive compounds [14, 15]. Individually, these compounds possess many useful bioactive properties, such as antibacterial [16, 17, 18], anti‐inflammatory [19], and antioxidant activity [20]. In combination, essential oil components have the potential to serve a multifunctional role by providing a coordinated suite of therapeutic activity that addresses diverse complications associated with implanted and indwelling devices. Among plant‐derived essential oils, peppermint essential oil is particularly notable for its high content of bioactive monoterpenoids, including menthol, menthone, and eucalyptol, which collectively confer broad biological properties, including antibacterial, antioxidant, anti‐inflammatory, and neuroprotective effects [21, 22, 23, 24]. Peppermint essential oil has been reported to reduce the expression of pro‐inflammatory cytokines such as IL‐6 and TNF‐α [25], helping to alleviate the inflammatory response [21]. Additionally, it has been demonstrated to enhance skin repair, promote fibroblast proliferation, and regulate TGF‐β signalling in vivo, improving wound healing in a murine model [26]. The use of peppermint essential oil as a diverse therapeutic agent is substantiated by both in vitro [25] and in vivo [26] evaluations.

The use of plant‐derived essential oils as coating precursors for medical devices has attracted increasing research attention in recent years. Current approaches typically involve the establishment of diffusion reservoirs that gradually leach the essential oil into the microenvironment surrounding the material [27, 28]. This approach suffers from intrinsic drawbacks such as a finite functional lifetime due to payload depletion, evaporative losses during fabrication [22], and degradation during storage [29]. Strategies to address these problems, such as encapsulation [30] and layer‐by‐layer assembly [31], can decelerate the release, but they often involve multi‐step and low‐throughput processes that are difficult to incorporate in large‐scale manufacturing. Furthermore, while these approaches may reduce diffusion rates, they do not fundamentally overcome the intrinsic limitation of a finite reservoir and, therefore, still suffer from a constrained functional lifespan.

A more attractive approach is to develop a contact‐active coating that retains its functionality over time. One report describes a technique of thermal polymerisation of peppermint essential oil [32], shifting the mode of action of the coating away from the typical diffusion reservoir, toward a coating that relies on interface‐driven activity. While this approach is a step in the right direction, current techniques still suffer from multiple major limitations. First and foremost, the thermal polymerisation approach described in that report selectively immobilizes only a subset of compounds in the peppermint essential oil, retaining in the final coating only 4 out of >20 starting compounds. This is a fundamental limitation of the thermal polymerisation process, which relies on specific chemical reactivity between components. Compounds that are highly volatile or unable to polymerise through solution‐based chemistry are excluded from the final coating. By losing the compositional identity of the parent precursor, the spectrum of functionality of the coating subsequently becomes narrower. This narrowing of functionality reduces the potential of the coating to holistically address the diverse complications associated with indwelling medical devices. A further limitation of the thermal polymerisation approach is its poor scalability, requiring chemical pre‐treatment of the substrate followed by a 48‐h thermal incubation period to facilitate polymerisation. Lastly, as polymerisation of the precursor molecules is governed by their chemical compatibility, the coating is less able to form densely crosslinked networks, minimizing coating stability.

In this report, we address these fundamental limitations by turning to plasma‐based deposition of peppermint essential oil, conducted at atmospheric pressure and room temperature. Plasma is often referred to as the fourth state of matter, consisting of a highly energetic gaseous mixture of charged molecules, ions, and electrons – and has been extensively used in material surface modification [33, 34]. The reactive environment of plasma enables the fragmentation and reassembly of precursor molecules into a highly crosslinked polymeric network on any substrate. A major advantage of plasma‐based polymerisation is that polymer coatings can be assembled from precursor molecules that could not typically be polymerised by conventional means [33]. In doing so, plasma‐based polymerisation enables the retention and incorporation of important functional groups from diverse bioactive precursors into a single, contiguous coating. This single‐step, solvent‐free process allows coatings to be applied to virtually any substrate, including complex 3D geometries. Further advantages of plasma‐based technologies are their conformity to green chemistry principles [35], non‐reliance on hazardous solvents, and a relatively low carbon footprint, which can be further reduced by harnessing renewable energy sources [36].

Our work advances the field of plant‐derived bioactive coatings by developing a facile one‐step approach to fabricate a multifunctional contact‐active coating via plasma‐polymerisation of peppermint essential oil (henceforth referred to as p(PEO)). While plasma‐polymerisation is known to be substrate independent, we aimed to pilot our approach using one application as a proof‐of‐concept model: silicon‐based urinary catheters. The use of urinary catheters as a model for this study is valuable for multiple reasons. First, the contact‐active antioxidant and anti‐inflammatory activity of p(PEO) are well positioned to alleviate the mechanically induced sterile inflammatory response that frequently occurs during bladder catheterisation. In parallel, the antibacterial activity of the coating is expected to minimize bacterial colonisation of the material, which is exceedingly common during catheterisation. To validate the bioactive multifunctionality of p(PEO), we evaluated its antioxidant, antibacterial, and immunomodulatory properties within the clinical context of bladder catheterisation. Our results represent a major advancement by confirming that a) plasma‐polymerisation of peppermint essential oil retains the diversity of bioactive functional groups present in the precursor oil and forms a stable coating; and b) the contact‐active coating inherits the multifunctionality of the individual components that comprise the essential oil, enabling it to reduce bacterial viability, scavenge reactive species and modulate the immune response to favour resolution of inflammation.

2. Results and Discussion

2.1. Primary Composition of Peppermint Essential Oil used in this Study

Prior to applying peppermint essential oil as a plasma‐deposited coating, we quantified its chemical composition using gas chromatography mass spectroscopy, resulting in the identification of 21 individual chemical components (Figure S1 and Table S1). From this full composition, we further identified that 6 components were present in concentrations above 1% (Table 1), while the remaining 15 components ranged from 0.03% to 0.75%. We expect that any bioactive properties measured in the plasma‐polymerised peppermint essential oil coating will be primarily attributable to the functional groups derived from the major constituents of the oil. Bioactivity of menthol (constituting 47.25% of the oil) is often attributed to both its hydroxyl functional group and its cyclohexane ring [37, 38]. Polar interactions driven by the hydroxyl group and hydrophobic interactions driven by the cyclohexane work in concert to promote antimicrobial activity via membrane disruption (especially in Gram‐negative bacteria) [38]. Similarly, these two functional components of menthol also contribute to immunomodulatory effects via multiple mechanisms [37]. The second most concentrated component of peppermint essential oil is L‐menthone (32.46%), which contains a ketone carbonyl group that has been reported to contribute to antibacterial activity via hydrogen bonding with phospholipid headgroups and subsequent interference with lipid packing in the envelope [39]. The ketone group of L‐menthone is also reported to contribute to antioxidant activity by participating in electron transfer with reactive species [39]. The presence of trans‐carane (6.34%) represents another contributor to antimicrobial activity, as its bicyclic and lipophilic structure enables membrane insertion and disruption [40]. Eucalyptol (5.54%) consists of a lipophilic bicyclic monoterpene structure with radical scavenging activity, and also drives multiple interactions that modulate the inflammatory response [41]. Caryophyllene (2.70%) has been reported to contribute to anti‐inflammatory activity by reducing the expression of LPS‐induced pro‐inflammatory cytokines [42]. The presence of D‐limonene (2.16%) is expected to contribute to anti‐oxidant activity due to its direct radical scavenging ability, but also indirectly via promoting cellular expression of antioxidant enzymes such as superoxide dismutase and catalase [43].

TABLE 1.

The most abundant compounds comprising peppermint essential oil used in this study. Cutoff was set at >1% of full composition.

Chemicals	Proportion (%)	Relevant biological activity	Refs.
Menthol	47.25	anti‐inflammatory, antibacterial, antifungal, and antiviral	[37, 38]
L‐Menthone	32.46	antioxidant, anti‐inflammatory, antimicrobial, and neuroprotective.	[39]
trans‐Carane	6.34	antibacterial activity	[40]
Eucalyptol	5.54	anti‐inflammatory, antioxidant, antimicrobial	[41]
Caryophyllene	2.70	anti‐inflammatory	[42]
D‐ Limonene	2.16	antioxidant, anti‐inflammatory, and anticancer	[43]

2.2. Preparation and Characterisation of the p(PEO) Coatings

Coatings were prepared using a custom‐built atmospheric pressure plasma deposition device, which is depicted in Figure 1A and described in the Materials and Methods section. Optical emission spectroscopy (OES) was utilized to investigate the properties of the plasma phase generated using pure argon gas and a mixture of argon gas and p(PEO) (Figure 1B). In both cases, the spectra revealed characteristic peaks of ionized argon (4p→ 4s) in the range of 700–900 nm [9, 44, 45]. These peaks confirm that the carrier gas was introduced successfully into the plasma system. The spectra also revealed peaks corresponding to ionized OH at 308 nm [44, 46] and N₂ molecules at 316 and 337 nm [44, 47]. The presence of these peaks is attributable to the composition of atmospheric gases. When the peppermint essential oil precursor was introduced, a broad range of new peaks appeared in the spectra, notably, the CH peak at 431 nm [48, 49] and C₂ (Swan band) at 516 and 563 nm [44, 50], which indicates the ionization of volatile compounds present in the oil. Furthermore, the change in the plasma color from violet (characteristic of pure argon gas) to light blue is simple visual qualitative evidence of the difference in the composition of the ionized gas inside the plasma reactor. The operating temperature of the plasma chamber was relatively low, reaching 36.5°C after 5 min, and peaking at 39.1°C after 10 min (Figure S2).

FIGURE 1.

Plasma‐deposited peppermint essential oil coatings were prepared using atmospheric pressure plasma deposition. (A) Schematic of the atmospheric pressure plasma system used for deposition. (B) The optical emission spectra of argon plasma alone and after introducing peppermint essential oil.

To optimize the plasma process parameters and maximize the thickness of the p(PEO) coating, we systematically evaluated the influence of argon gas flow rate, working power, and process duration (Figure 2A–C). A thickness of 31 nm (measured by spectroscopic ellipsometry) was achieved using a 3 L min⁻¹ gas flow rate, 5 kV working power, and 10 min process duration. Having established this, we aimed to evaluate the influence of coating thickness on the physical and chemical properties of the coating, as well as its biological performance. To do so, we utilized two different coating durations, 5 and 10 min (resulting in coatings of ≃ 17 and ≃ 31 nm thickness, respectively). In order to further validate the deposition under these parameters, we employed a masking technique to prepare substrates containing both coated and uncoated regions, then used scanning electron microscopy (SEM) to visualize the coatings (Figure S3). The SEM micrographs revealed clearly distinct coated and uncoated regions, providing microscopic validation of a successful deposition process.

FIGURE 2.

Optimization and characterization of the p(PEO) coatings. (A) Influence of argon gas flow rate on coating thickness. Working power fixed at 2.5 kV for 1 min; (B) Influence of working power on coating thickness. Argon gas flow rate fixed at 3 L min⁻¹ for 1 min (C) Influence of coating duration on coating thickness. Working power fixed at 5 kV and argon gas flow rate fixed at 3 L min⁻¹; (D,E) Reduction in thickness coating after 24 h exposure to deionized water and PBS, respectively; (F) ζ‐potential of 10‐min coating. (G,H) AFM micrographs showing the topography of 5 and 10‐min coatings, respectively. The scale bar displays 1 µm (I) Water contact angle; (J) FTIR of 5 and 10‐min coatings. (K,M) XPS survey spectra of 5‐ and 10‐min coatings, respectively; (L,N) C1s high resolution spectra of 5‐ and 10‐min coatings, respectively. All experiments were performed in triplicate. Data were analysed using a one‐way ANOVA, using Bonferroni post hoc analysis for multiple comparisons. Data plotted as mean ± SD, n = 3, ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001, and ns = not significant.

Next, we examined the coating stability by immersing them in either Milli‐Q water or PBS (Figure 2D,E) for 24 h and then measured the reduction in coating thickness. We found that the coatings were highly stable in these physiologically relevant media. The thickness of the coatings was reduced by only 2% and 4%, respectively. We also evaluated the surface charge of the p(PEO) coating by measuring its zeta (ζ)‐potential in the pH range between 3 and 9 (Figure 2F). The coatings were negatively charged, with the ζ‐potential decreasing from −5 to −15 mV with increasing pH. At physiological pH = 7, the coatings had a charge of −11.23 ± 0.89 mV.

In order to characterize the topographic features of p(PEO), we utilized atomic force microscopy (Figure 2G,H). The coating deposited for 5‐min had Root Mean Square roughness (RMS) of 6.2 nm and displayed larger topographical features. The coating deposited for 10 min was significantly smoother with an RMS of 1.6 nm. The decrease in roughness at longer deposition durations suggests that the mechanism of coating growth from the surface may involve the initial formation of discrete, ‘island‐like’ nuclei which gradually coalesce as more polymer material is deposited, shifting the surface topography from a rough landscape with abundant peaks and troughs, toward a smooth coating devoid of peaks. This interpretation is supported by previous research that mapped the ‘island‐like’ formation pathway of plasma polymers derived from n‐heptylamine [51].

We measured the wettability of the coating by the sessile drop water contact angle (WCA; Figure 2I). The uncoated silicon substrate had a WCA of 48.4° ± 1.3°. The 5‐ and 10‐min coatings both had a significantly higher WCA (p < 0.05) in comparison to the uncoated silicon substrate (73.4° ± 2.7° and 81.1° ± 5.5°, respectively), but the difference between the two coatings was not statistically significant. To investigate the chemical structure of p(PEO), we first utilized Fourier‐transform infrared (FTIR) spectroscopy to examine the chemical bonds present in the coatings, as well as the raw oil precursor (Figure 2J). The resulting spectra showed that the 5‐ and 10‐min coatings had a highly similar profile, and these were distinct from the raw oil, with most differences identified in the 1500–1000 cm⁻¹ (fingerprint) region. Specifically, the raw oil had multiple distinct peaks at 1563 cm⁻¹ (aromatic stretching) [52], 1453 cm⁻¹ (C─H bending) [53], 1332 cm⁻¹ (aromatic ring) [54], 1205 cm⁻¹ (C─O─C stretching, ethers) [55], and 1053 cm⁻¹ (C─O stretching, primary alcohols) [56]. All of these chemical signatures are attributable to the various structural constituents of the peppermint essential oil (listed in Table S2). In contrast, in the 1500–1000 cm⁻¹ region of the FTIR spectra generated for 5‐ and 10‐min coatings, the spectra displayed one large, broad peak that encompassed all individual peaks associated with the raw oil. This spectral profile is consistent with a loss of the discrete molecular structures within the precursor essential oil components, and formation of an amorphous and highly crosslinked polymeric structure resulting from plasma deposition. Additionally, we noted that the 10‐min coating had a distinct peak at 1699 cm⁻¹, which we attributed to conjugated C═O bonds [57], indicating formation of additional carbonyl functionalities. We further supported this analysis using X‐ray photoelectron spectroscopy (XPS), Figure 2K–M. As expected, p(PEO) surfaces exhibited peaks associated with O1s and C1s. We analyzed the high‐resolution spectra in the C1s region to obtain further insight into the chemical state of carbon‐associated bonds. Echoing the findings of the FTIR analysis, we identified components attributable to aliphatic hydrocarbons (C─C, C─H) at 285 eV [58, 59] and the C─OH or C─O─C components at 286.2 eV [59, 60]. The presence of these bonds can be traced back to the peppermint oil precursor composition, which includes compounds such as menthone and methyl acetate that exhibit ketone or ester linkages (Table S1 and Figure S1). The peak at 287.5 eV corresponding to C═O indicates the presence of carbonyl or carboxyl functional groups [59, 61]. The detected O─C═O components at 288.5 eV [59, 60] further implicate the presence of carboxyl functional groups, which are likely the source of the negative charge of the coatings discussed above. Overall, the chemical analyses confirmed the successful deposition of polymerized peppermint oil and verified the absence of contaminants. In addition to this, we considered the possibility that reactive species generated in the plasma chamber could potentially remain on the coating surface, which may influence biological cells in subsequent analyses. To investigate this, we used a ROS detection kit to measure the presence of reactive species 20 min after coating deposition, as well as 24 h later (Figure S4). In the first 20 min after plasma exposure, we detected a significant presence of reactive species on samples treated with argon plasma alone or with peppermint essential oil. However, after repeated measurements of the same samples at 24 h, all plasma‐treated groups displayed a considerably lower presence of reactive species, equal to the group not exposed to plasma. This behaviour is consistent with the inherently short lifetimes of plasma‐generated reactive species and indicates that any transient oxidative effect from the coating would dissipate well before biological exposure.

2.3. Antioxidant Activity of the p(PEO) Coatings

To evaluate the antioxidant activity of p(PEO) coatings, we utilized a radical scavenging assay, which measures the conversion of the radical form of 2,2‐diphenyl‐1‐picrylhydrazyl (DPPH•, violet in colour) to DPPH‐H (colourless) in the presence of antioxidants. p(PEO) coatings were deposited on a tissue culture well plate and a silicon urinary catheter tube (Figure 3A), the latter serving as a model biomedical device. When the tissue culture plate was coated for 5 min, a 3 h incubation with DPPH resulted in 21.63 ± 2.31% scavenging of initial free radicals, which increased to 31.67 ± 2.08% after 6 h of incubation (Figure 3B). A comparable trend was noted with 10‐min coatings, which scavenged 22.27 ± 2.08% of initial DPPH radicals after 3 h and increased to 32.67 ± 2.11% after 6 h exposure. The coated samples had superior DPPH radical scavenging activity (p < 0.05) compared to the untreated samples, which exhibited values of 3.34 ± 1.53% after 3 h of incubation and 8.71 ± 1.5% after 6 h of experiment. When we coated p(PEO) on the silicon urinary catheter, we observed a similar but more pronounced trend, resulting in approximately 25% and 40% radical scavenging with 3 and 6 h incubation times, respectively (Figure 3C). Compared to the coated tissue culture plate, the heightened radical scavenging ability of the catheter may be due to the increased surface area to volume ratio of the catheter geometry. To support and validate our DPPH assay, we employed a second evaluation of antioxidant activity using (2,2'‐azino‐bis(3‐ethylbenzothiazoline‐6‐sulfonic acid) (ABTS) as the reagent. Again, we tested both a p(PEO) coated tissue culture plate and a urinary catheter. The results showed a similar trend to the DPPH assay but were more pronounced. For the DPPH assay, the highest antioxidant percentage was approximately 40% (6 h incubation with the catheter coating), whereas for the ABTS⁺ assay, the highest measured antioxidant activity was approximately 90% with the same conditions. This disparity may be due to a combination of factors, including structural differences between ABTS and DPPH (influencing steric interactions between reagent and surface‐bound reductive groups present in the polymer matrix), and differences in chemical reactivity. The ABTS radical exhibits reactivity with a broader spectrum of antioxidants, including some that do not respond effectively to DPPH. For example, it was reported that dihydrochalcones and flavanones do not react with DPPH but do react with ABTS [62, 63]. Taken together, these results suggest that multiple functional moieties within the p(PEO) matrix contribute to its robust antioxidant activity, and that the measured effects arise from true redox interactions at the coating–solution interface rather than analytical artefacts.

FIGURE 3.

Free radical scavenging ability of the coating against DPPH and ABTS⁺ radicals. (A) Experimental procedure; (B) The radical scavenging ability of the p(PEO) coatings deposited on the tissue culture well plate; (C) The radical scavenging ability of the p(PEO) coatings deposited on a catheter tube. All experiments were performed in triplicate. Data were analysed using a one‐way ANOVA, using Bonferroni post hoc analysis for multiple comparisons. Data plotted as mean ± SD, n = 3, ^* p < 0.05, and ns = not significant.

2.4. Antibacterial Activity of the p(PEO) Coatings

We applied coatings to flat silicon wafers to assess the antibacterial effectiveness of p(PEO) coatings. Our method included Live/Dead fluorescence analyses alongside traditional colony enumeration, against two clinically relevant Gram‐positive (S. aureus and S. epidermidis) and two Gram‐negative (E. coli and P. aeruginosa) pathogens. As a general observation, we noted that the p(PEO) coating was significantly more effective against Gram‐negative bacteria than Gram‐positive bacteria, under the conditions we tested. For P. aeruginosa, the Live/Dead confocal micrographs showed that a 6 h incubation on p(PEO) coatings resulted in 85.1% and 89.4% red‐stained cells, for coatings deposited for 5 and 10 min, respectively, which slightly decreased to 75.2% and 82.4% after 24h of incubation (Figure 4A). The difference in the percentage of red‐stained cells at 6 and 24 h of incubation indicates that the bactericidal properties of the p(PEO) coatings are most effective during the initial hours of cell attachment, and that longer incubation times do not further increase the effect. The Live/Dead analyses were supported by colony enumeration data (Figure 4B), in which the 5 and 10‐min coatings had 8.36×10⁵ and 6.43×10⁵ CFU/mL, respectively, compared to the 1.25×10⁶ CFU/mL of the non‐treated substrate (after a 6 h incubation). Similarly, for E. coli, a 6 h incubation with p(PEO) resulted in 72.5% and 79.1% red‐stained cells for 5 and 10‐min coatings, respectively. This was supported by the colony enumeration results, which showed that a 6 h incubation resulted in approximately 1.84 × 10⁶ and 1.07 × 10⁶ CFU/mL, for 5‐min and 10‐min coatings, respectively, reduced from the 2.57 × 10⁶ CFU/mL on the non‐treated sample. These values represent a moderate antibacterial activity driven by the p(PEO) coating, contributing to an approximate 50%–70% reduction in viable cell counts compared to the uncoated substrate. This degree of activity can be compared to another form of peppermint oil coating produced by a different laboratory [32]. In their report, the group described a thermally polymerised peppermint oil coating on titanium substrates. In that report, the authors measured an antibacterial activity of approximately 85%. The moderately higher antibacterial activity in that report could be attributed to the instability of thermally polymerised peppermint oil; this is substantiated within the report, showing that the coating is gradually washed off. This leaching of bioactive components from the coating may enable the free‐floating components to interfere with membrane integrity. Nevertheless, this moderate increase in antibacterial activity comes at a cost of a weaker durability and shorter functional lifespan.

FIGURE 4.

Antimicrobial activity of the p(PEO) coatings against Gram‐negative and Gram‐positive bacteria. (A) Confocal microscopy images of P. aeruginosa and E. coli stained using the Live/Dead assay. Scale bars represent 20 µm. (B) Quantitative colony counts of P. aeruginosa and E. coli. (C) Confocal microscopy images of S. aureus and S. epidermidis stained using the Live/Dead assay. Scale bars represent 20 µm. (D) Quantitative colony counts of S. aureus and S. epidermidis. All experiments were performed in triplicate. Data were analysed using a one‐way ANOVA, using Bonferroni post hoc analysis for multiple comparisons. Data plotted as mean ± SD, n = 3, ^* p < 0.05, ^** p < 0.01, ^**** p < 0.0001 and ns = not significant.

In our results, we note a disparity in the percentage of red‐stained cells (75%–90%) compared to the reduction in CFU counts between coated and non‐coated samples (30%–70%). This disparity is likely due to the different principles on which these two assays operate. Specifically for the Live/Dead assay, propidium iodide (PI; red stain) is taken up by cells with a compromised cell envelope, and the presence of PI fluorescence is used as a proxy for quantifying cell death. However, some cells with disturbed envelopes may remain viable, proceeding to recover and divide. This phenomenon of sub‐lethal injury followed by recovery has been reported to occur in relevant literature. Specifically, this has been observed in relation to varying stress triggers, including acidic/alkaline shock, thermal, physical, and osmotic stresses [64, 65]. The effects of sub‐lethal injury include structural damage to the cell envelope, thereby permeabilizing the cells to external compounds. Thus, it is likely that the bactericidal percentages captured by our Live/Dead analyses also include sub‐lethally injured cells that have internalised PI but recover and remain viable. The colony enumeration data, therefore, provides a more accurate indicator of the actual viability of cells. Nevertheless, the information gained from our Live/Dead analysis is still important, because it suggests that the p(PEO) coating not only kills cells but also permeabilizes the membrane of a subset of surviving cells; thus potentially offering a window of opportunity for the immune system or a combination therapy approach to eliminate the remaining cells.

To better understand the mechanism of antibacterial action, we performed a zone of inhibition analysis (Figure S5) to evaluate whether the antibacterial effect was contact‐dependent or based on diffusion of bioactive compounds. We observed that bacterial growth was only inhibited when in direct contact with the coated substrates, and no zone of inhibition was recorded around the substrates–indicating a contact‐killing effect. This therefore suggests that the bactericidal activity of the p(PEO) coatings occurs exclusively at the material interface. Relevant literature attributes the antibacterial activity of peppermint essential oil to its terpenoid‐rich composition [66], which can disrupt the bacterial membrane. This is commensurate with our Live/Dead analysis, which shows a high degree of membrane permeabilization.

In contrast to the positive antibacterial results observed for Gram‐negative pathogens, there was a lack of bactericidal activity against Gram‐positive bacteria (Figure 4C,D). To shed further light on the antibacterial properties of the p(PEO) coatings, we measured membrane polarization using the BacLight Bacterial Membrane Potential kit (Figure S6). In this assay, a loss of membrane polarization is indicated by a reduction in the red/green fluorescence ratio. We observed that, when P. aeruginosa was cultured on the coated substrates, the red/green ratio was reduced from 0.9 (non‐coated substrate) to 0.3 (5 and 10‐min coatings), performing similarly to the membrane‐depolarizing control (CCCP; red/green ratio of 0.25). In contrast, when S. aureus was investigated, the red/green ratio was similar between the non‐coated and coated substrates. Bacterial cells rely on membrane polarization to maintain protonmotive gradients and critical metabolic processes [67]. Therefore, the membrane depolarizing activity of the p(PEO) coating likely plays a key role in its antibacterial performance. The variation in coating performance against Gram‐positive and Gram‐negative bacteria likely reflects fundamental differences in their cell wall architecture. The peptidoglycan layer of Gram‐positive bacteria ranges from 20 to 80 nm in thickness and encapsulates the cytoplasmic membrane. In Gram‐negative bacteria, the peptidoglycan layer is significantly thinner (1.5–10 nm) and is nestled between two membrane bilayers [68]. As the bactericidal activity of phenolic compounds is reported to be driven by membrane‐disruptive effects, the thick peptidoglycan layer of Gram‐positive bacteria likely explains the lack of activity against these species. Specifically, the thick peptidoglycan of Gram‐positive bacteria likely acts as a barrier that shields the cytoplasmic membrane from the bioactive functional groups in the p(PEO) coatings.

2.5. Antibacterial Functionality of the p(PEO) Coatings after Immersion in PBS

As discussed above, the p(PEO) coatings were stable in PBS for 24 h, but it is also important to investigate whether the activity of the coatings was retained following this period of exposure (Figure 5A). To do so, we immersed the p(PEO) coatings in PBS for 24 h, and repeated the antibacterial assays previously described, using P. aeruginosa and E. coli as the tested pathogens. The Live/Dead confocal micrographs and corresponding percentages of red‐stained cells (Figure 5B,D) demonstrated that the coatings retained their efficacy against these Gram‐negative pathogens. The 5‐min coating resulted in 70% red‐stained cells when incubated with P. aeruginosa and 68% when incubated with E. coli, whereas the 10‐min coating achieved 80% and 78% dead cells, respectively. These fluorescence‐based measurements were again supported by colony enumeration (Figure 5C). Viable cell counts of P. aeruginosa were 1.88 × 10⁷ and 1.31 × 10⁷ CFU/mL when cultured on 5 and 10‐min coatings, respectively. This represents a reduction from the number of viable cells recovered from non‐coated samples, which was 2.97 × 10⁷ CFU/mL. The results were nearly identical for E. coli, with which we observed 3.79 × 10⁷ viable cells on the non‐coated samples, which was reduced to 1.97 × 10⁷ and 1.73 × 10⁷ CFU/mL for 5‐ and 10‐min coatings, respectively. These results demonstrate that the coatings retain their bactericidal activity even after pre‐immersion in PBS for 24 h.

FIGURE 5.

Longevity of the coating against Gram‐negative bacteria. (A) Experimental procedure. (B) Confocal microscopy images of P. aeruginosa and E. coli stained using the Live/Dead assay. Scale bar represents 30 µm. (C) Quantitative colony counts of P. aeruginosa and E. coli. (D) Qualitative analysis of the confocal microscopy images of P. aeruginosa and E. coli stained using the Live/Dead assay. All experiments were performed in triplicate. Data were analysed using a one‐way ANOVA, using Bonferroni post hoc analysis for multiple comparisons. Data plotted as mean ± SD, n = 3 and ^* p < 0.05, ^** p < 0.01, ^**** p < 0.0001.

2.6. Antibacterial Activity of p(PEO)‐Coated Urinary Catheters

Catheter‐associated urinary tract infections are among the most common hospital‐acquired infections, with E. coli and P. aeruginosa representing the predominant causative pathogens. Thus, the selective efficacy of p(PEO) coatings against Gram‐negative bacteria aligns with a significant unmet need to prevent catheter‐associated infections [69]. Hence, in the next step, we prepared p(PEO)‐coated silicon urinary catheters (Figure 6A) and tested their antibacterial efficacy against multiple Gram‐negative bacteria, including a clinical isolate of a uropathogenic E. coli (UPEC) (Figure 6B). The latter was important since clinical pathogen isolates are genotypically distinct from standard laboratory strains, with unique structural and virulence characteristics. The results aligned closely with our antibacterial evaluations discussed above. When P. aeruginosa was used, the number of viable cells was 1.59 × 10⁷ and 1.27 × 10⁷ CFU/mL for the 5 and 10‐min coatings, respectively, which was reduced from the non‐treated catheter that showed 2.27 × 10⁷ CFU /mL. Similarly, for the laboratory strain of E. coli, 5‐ and 10‐min coatings had 1.70 × 10⁷ and 1.57 × 10⁷ CFU/mL, while the non‐coated catheter had 2.55 × 10⁷ CFU/mL. Lastly, for the clinical UPEC isolate, the non‐coated catheter had 1.85 × 10⁷, while the 5‐ and 10‐min coatings reduced the viable cell counts to 1.08 × 10⁷ and 9.03 × 10⁶ CFU/mL. These results indicate that the geometry and bulk properties of the coated substrate have no significant influence on the antibacterial properties of the coating, as the modified catheter material performs equally as effectively as when the coatings were applied on a flat silicon substrate. Second, this is validation that the p(PEO) coatings retained their antibacterial potency against a clinical UPEC isolate despite the unique structural and virulent characteristics of these pathogens.

FIGURE 6.

The activity of p(PEO) coatings deposited on a urinary catheter, against Gram‐negative pathogens and a clinical isolate. (A) Experimental workflow. (B) Quantitative colony counts of P. aeruginosa, E. coli, and clinical isolate of uropathogenic E. coli (C). SEM images of P. aeruginosa, E. coli, and clinical isolate E. coli cultured with the p(PEO) coating deposited for 10 min. Yellow arrows denote cells exhibiting morphological abnormalities. The scale bar, bottom right image, represents 5 µm. All experiments were performed in triplicate. Data were analysed using a one‐way ANOVA, using Bonferroni post hoc analysis for multiple comparisons. Data plotted as mean ± SD, n = 3, ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001, and ns = not significant.

To qualitatively evaluate the effect of the p(PEO) coatings on the morphology of the studied pathogens, we analyzed them at high magnification using SEM (Figure 6C). The cells appeared to have their typical morphology on the non‐coated catheter material, displaying smooth outer surfaces and a turgid structure. In contrast, the cells incubated on the p(PEO) coatings often appeared flattened, lysed, or otherwise disturbed from their typical morphology. Cells with these characteristics are highlighted with yellow arrows.

2.7. The p(PEO) Coatings Enhanced the Activity of Clinically Relevant Antibiotics–A Combination Therapy Approach

Our results demonstrated that incubating Gram‐negative bacteria on p(PEO) coatings led to a substantial proportion of PI‐permeable cells, indicating either loss of viability or sub‐lethal injury. We hypothesized that these sub‐lethally injured populations would exhibit increased susceptibility to subsequent antibiotic treatment. This hypothesis is substantiated by diverse literature that proposes multiple pathways that lead to enhanced antibiotic activity. Membrane permeabilizers [70], for example, have been reported to promote the influx of antibiotics which may otherwise be bottlenecked by the membrane barrier. This concept is specifically relevant to essential oils [71] and phenolic compounds [72]. On top of promoting influx, it is also believed that essential oils can minimize efflux, either by blocking transporters or by interfering with membrane polarity. Interestingly, one study reported that menthol may inhibit efflux pumps in Acinetobacter baumannii, thereby significantly reducing minimum inhibitory concentrations of ciprofloxacin [73]. While most antibiotic potentiation literature is focused on solution‐based combination therapies, there is emerging evidence that surface‐mediated stresses can promote the activity of antibiotics [74, 75]. In relation to this, there is a growing understanding of the importance of material surface properties in the context of antibiotic activity against surface‐attached cells [76]. In a clinical context, it is common for catheterized patients to receive prophylactically administered antibiotic treatments to minimize the occurrence of urinary tract infections (UTIs) [77]. Based on this, we determined whether E. coli (including both the laboratory strain and the UPEC isolate) would become sensitized to colistin or levofloxacin following incubation on p(PEO) coatings. These antibiotics were chosen specifically for their relevance in treating and preventing UTIs [77, 78]. To simulate a prophylactic treatment protocol, we first incubated the pathogens on the p(PEO)‐coated catheters for 15 min and 3 h, then dosed the culture media with either levofloxacin or colistin (Figure 7A) at a concentration equivalent to the minimum bactericidal concentration determined for planktonic cultures (Figure S7). As a general trend across all groups, when antibiotic treatment was applied to bacteria attached to the non‐coated catheter, the viable cell counts were reduced by approximately 40%–70%. It should be noted that this degree of bactericidal activity was observed even when cells were only attached for 15 min prior to antibiotic treatment, reflecting the rapid change in phenotype that bacterial pathogens undergo when attaching to a surface. When the pre‐treatment attachment duration was extended to 3 h, the viable cell counts increased across all treatment types, indicating that the attached cells gradually gained higher antibiotic tolerance. Encouragingly, antibiotic treatment of cells incubated on the p(PEO)‐coated catheters resulted in significantly fewer viable cells compared to cells attached to non‐coated catheters. We note that this effect was independent of the coating duration, as there was no significant difference in viable counts between 5 and 10‐min coated surfaces. When colistin was applied after 15 min of cell attachment, the pattern of activity was almost identical between the laboratory strain and clinical isolate, reducing the viable counts from approximately 2.9 × 10³ CFU/mL on the non‐coated catheter to approximately 1.6 × 10³ CFU/mL on the p(PEO)‐coated catheter (Figure 7B). With 3 h of pre‐treatment attachment, there was a slightly different response between the laboratory E. coli strain and the clinical UPEC isolate. In this case, the quantity of viable cells of the laboratory strain on the colistin‐treated non‐coated catheter was 7.2 × 10³ CFU/mL, while the colistin‐treated p(PEO) catheters had approximately 3.2 × 10³ CFU/mL. Under the same conditions, the UPEC isolate showed 6.4 × 10³ CFU/mL on the non‐coated catheter, while the p(PEO) catheters showed approximately 3 × 10³ CFU/mL (Figure 7D). An overall similar trend was observed when levofloxacin was applied. In this case, the laboratory strain and clinical UPEC isolate responded almost identically to treatment. When the cells were given 15 min to attach before treatment, the levofloxacin‐treated non‐coated catheter had 1.4 × 10⁴ CFU/mL. At the same time, the p(PEO) coatings yielded approximately 7.6 × 10³ CFU/mL and 4.6 × 10³ CFU/mL for 5 and 10‐min coatings, respectively (Figure 7C). When the pre‐treatment attachment duration was increased to 3 h, the levofloxacin‐treated non‐coated catheter had approximately 3.4 × 10⁴ CFU/mL, while the p(PEO) coatings had 1.7 × 10⁴ and 1.3 × 10⁴ CFU/mL, for 5 and 10‐min coatings, respectively (Figure 7E).

FIGURE 7.

Coating‐mediated potentiation of clinically relevant antibiotics. (A) Experimental procedure. (B) Viable cell counts of E. coli (laboratory and clinical isolates) exposed to colistin, when pre‐treatment attachment was 15 min. (C) Viable cell counts after exposure to levofloxacin, when pre‐treatment attachment was 15 min. (D) Viable cell counts after exposure to colistin, when pre‐treatment attachment was 3 h. (E) Viable cell counts after exposure to levofloxacin, when pre‐treatment attachment was 3 h. All experiments were performed in triplicate. Data were analysed using a one‐way ANOVA, using Bonferroni post hoc analysis for multiple comparisons. Data plotted as mean ± SD, n = 3, and ^* p < 0.05.

Overall, these data show a consistent trend where the activity of clinically relevant antibiotics improves when interacting with Gram‐negative pathogens incubated on the p(PEO) coated surfaces. It is worth noting that the peppermint essential oil precursor we used contained 47.25% menthol, which has been reported to interfere with drug efflux in A. baumannii [73]. Our results further advance this concept, showing that antibiotic potentiation can occur even when the relevant molecular functionalities are part of a broad, immobile polymer network. This bodes well for clinical translation of the coating, because it benefits from the well‐established standard procedure of antibiotic prophylaxis that is commonly applied prior to bladder catheterization. Furthermore, by combining the bactericidal activity of the p(PEO) coating with conventional antibiotics, the combined outcome involves antibacterial action stemming from multiple mechanisms, which may assist in mitigating the emergence of antimicrobial resistance [79]. In principle, this is due to the way in which antibiotic resistance emerges. When cells with low‐level antibiotic resistance can survive conventional antibiotic regimes, they act as a gene pool to fuel the proliferation of a subsequent generation of resistant cells, which continues through a process of selection [80]. However, the presence of an alternative mechanism of antibacterial activity can eliminate those early‐generation resistant cells before they establish a highly resistant population.

2.8. Cytocompatibility of the p(PEO) Coating

A prerequisite of any biomaterial is its ability to interface with mammalian cells without triggering detrimental effects. To evaluate the biocompatibility of the p(PEO) coating, we conducted qualitative morphological assessments using actin/DAPI staining of HaCaT cells and differentiated THP‐1 cells, and used the MTT assay to measure metabolic activity (Figure 8A). In addition, we also measured intracellular ROS to evaluate whether the coating triggers oxidative stress. Qualitative analysis of the actin/DAPI fluorescence confocal micrographs (Figure 8B) shows comparable cell density and morphology across the groups cultured on the non‐coated substrates and the p(PEO) coatings, indicating that the structural properties and proliferation of both cell types are unhindered by the coating. Supporting this, the MTT assay (Figure 8D,E) yielded viability between 90% and 100% normalized to TCP, indicating that the metabolism of HaCaT and THP‐1 macrophage‐like cells is completely unhindered over a 72‐h period. Measurement of fluorescence intensity following DCF staining in THP‐1 cells (Figure 8C,F) showed that the p(PEO) coatings do not induce elevated levels of ROS within the cells. Conversely, the fluorescence was more intense in the cells attached to the non‐coated surface. This is an encouraging observation, because it affirms that the p(PEO) coatings eliminate extracellular ROS (as demonstrated in Figure 3) and minimize intracellular ROS in attached macrophages. We also performed a haemolysis assay to evaluate the blood compatibility of the p(PEO) coating (Figure S8). All coated substrates had <0.5% haemolytic activity, normalized to a Triton X control. This evaluation is important because it suggests that the p(PEO) coating could feasibly be extended to blood‐contacting applications, such as central venous catheters. Collectively, these results represent in vitro validation of the biocompatibility of the p(PEO) coatings by the absence of negative influence on the morphology, proliferation, or metabolic activity of mammalian cells, and overall, promote a favourable environment for host tissue vitality. These results are consistent with other reports demonstrating the cytocompatibility of plasma‐polymerised essential oils, including a coating derived from oregano oil [81], which showed no cytotoxic effects toward dermal fibroblasts. Although the oregano‐oil plasma polymer is formed from a precursor with a markedly different chemical composition from peppermint oil, the absence of cytotoxicity in both systems provides broader validation that plasma polymerisation of naturally derived oils can yield stable, biocompatible coatings.

FIGURE 8.

Biocompatibility analysis of p(PEO) coating. (A) Experiment workflow of MTT assay on differentiated THP1 cells and HaCaT cells. (B) Confocal micrographs of differentiated THP1 macrophage cells and HaCaT cells stained with phalloidin and DAPI. Scale bars represent 20 µm. (C) Confocal micrographs of THP1 cells stained with DCF to measure ROS presence. Scale bars represent 20 µm. (D) Quantified cell viability of HaCaT cells, based on MTT analysis, normalized to tissue culture plate (TCP). (E) Quantified cell viability of THP‐1 macrophages, based on MTT analysis, normalized to tissue culture plate (TCP). (F) Quantified fluorescence intensity of DCF. Data plotted as mean ± SD, n = 3, and ns = not significant.

2.9. Immunomodulatory Properties of the p(PEO) Coatings

To evaluate the potential immunomodulatory properties of p(PEO) coatings, we cultured differentiated THP‐1 cells on the modified substrate. We employed an array of immunological assessments, including ELISA, quantitative gene expression, and fluorescence‐based analyses of CD206, a marker for M2 macrophages (Figure 9A). Furthermore, we evaluated the immune response in the presence and absence of endotoxin (lipopolysaccharide; LPS), simulating both aseptic inflammation as well as inflammation associated with bacterial colonization [82]. This is an important distinction, because it has been reported that catheterization triggers an inflammatory response almost immediately after insertion [83], which may lead to further complications. Initially, we used ELISA to quantify IL‐6 (pro‐inflammatory) and IL‐4 (anti‐inflammatory) cytokines (Figure 9B). For IL‐6, we noted an approximate 50% reduction on the p(PEO) coated surfaces compared to a non‐treated control. This was the case for both LPS‐stimulated and non‐LPS groups. Conversely, for IL‐4, the p(PEO) coating elicited a 40%–50% increase in cytokine concentration. To support these findings, we conducted quantitative gene expression analyses on two pro‐inflammatory cytokines (IL‐6 and TNF‐α) alongside two anti‐inflammatory cytokines (IL‐4 and IL‐10) (Figure 9C). The IL‐6 expression results were comparable to the corresponding ELISA measurements, with the non‐coated surface displaying a significantly higher expression compared to the p(PEO) coating (p < 0.05), irrespective of LPS stimulation. For TNF‐α, the relative normalized expression was approximately 1.25 on the non‐treated surface stimulated with LPS, compared to 1.0 for the p(PEO) under the same conditions (P < 0.05). For the anti‐inflammatory IL‐4, the p(PEO) coatings elicited an increase from approximately 0.6 to 0.8 relative normalized expression (p < 0.05), representing a moderate upregulation consistent with the ELISA assay. However, IL‐10 expression was not significantly different between coated and non‐coated surfaces. As IL‐4 and IL‐10 are both involved in the resolution of the inflammatory response, it initially seems counterintuitive that the p(PEO) coating only triggers upregulation in IL‐4 and not IL‐10. However, relevant literature has demonstrated that a reduced presence of ROS results in decreased production of IL‐10 [84]. We have demonstrated that the p(PEO) coating has ROS‐scavenging activity, likely contributing to quenching the IL‐10 production that ordinarily accompanies an anti‐inflammatory response.

FIGURE 9.

The immunomodulatory properties of p(PEO). (A) Experimental procedure. (B) ELISA assay of IL‐6 and IL‐4 cytokines. (C) mRNA expression of pro‐inflammatory and anti‐inflammatory cytokines from THP‐1 macrophages in the presence and absence of LPS. (D) Confocal micrographs showing the presence of CD80 and CD206. (E) Flow cytometry results of the proportion of macrophages positive for CD206. All experiments were performed in triplicate. Data were analysed using a one‐way ANOVA, using Bonferroni post hoc analysis for multiple comparisons. Data plotted as mean ± SD, n = 3, ^* p < 0.05, and ns = not significant.

Fluorescence‐based analyses of macrophage polarization were performed to assess the immunomodulatory effects of p(PEO) by quantifying the M1/M2 macrophage ratio. First, we captured confocal micrographs showing the presence of the pro‐inflammatory biomarker (CD80) and anti‐inflammatory marker (CD206) (Figure 9D). CD80 is a co‐stimulatory molecule primarily expressed on antigen‐presenting cells (APCs) like macrophages. Conversely, CD206 is a marker for alternatively activated macrophages (M2). The CD206/CD80 ratio measures macrophage polarization, with a higher CD206/CD80 ratio indicating a shift toward the anti‐inflammatory (M2) phenotype and a lower ratio indicating the pro‐inflammatory (M1) phenotype. Specifically, we observed that the ratio of CD206/CD80 in cells on the p(PEO)‐coated surfaces was approximately 1.1, more than 2‐fold higher than that of non‐coated surfaces, with a ratio of 0.5. This indicates that the proportion of M2 cells is significantly higher on the p(PEO) surface. A similar trend repeated when LPS was administered, as the signals of CD206 on the coating were greater than those of the non‐coated samples. Furthermore, we utilized flow cytometry to quantify the presence of M2 cells using CD206 as a biomarker (Figure 9E). The measurements indicated that the proportion of M2 cells in the population under unstimulated conditions was 6.1% and 6.3% for the 5 and 10‐min coating treatments, respectively, approximately 2‐fold higher than the proportion recorded in the non‐coated group, which exhibited 3.1% M2 cells. Under the influence of LPS, the percentage of M2 cells on the non‐coated substrate was 1.8%, whereas the quantities on the coated surfaces were 4.09% and 5.98%, for 5 and 10‐min coatings, respectively. Overall, these results all indicate that the p(PEO) coating triggers an immunological response that favours the resolution of inflammation, as pro‐inflammatory cytokines were downregulated, and IL‐4 was upregulated. The CD206 marker for M2 macrophages became more abundant. These observations align with the findings of relevant publications that report on the anti‐inflammatory activity of peppermint essential oil [25, 26].

A potential understanding of the underlying mechanism of immunomodulation can be gained by drawing upon the available relevant literature. In one insightful study, a group performed molecular docking studies to investigate the specific interactions that underpin the anti‐inflammatory effect of green cardamom essential oil (GCEO) [85]. Among multiple interactions, they found that a particular metabolite in GCEO, 1,8‐cineole (also known as eucalyptol), had a strong affinity for membrane‐bound protein tyrosine kinase 2B (PTK2B) in immune cells. The authors proposed this interaction as a key mechanism underlying the anti‐inflammatory effects of GCEO. This is highly plausible, as other related research has reported the role of PTK2B in the inflammatory response [86]. Our GC–MS results identified eucalyptol as the fourth most concentrated compound in the peppermint essential oil precursor, accounting for over 5% of the composition (Table 1). Given that plasma‐polymerization is known to produce polymers that retain many of the functionalities of the precursor, it is plausible that the p(PEO) coating inherits the ability of eucalyptol to dock with PTK2B, thus enabling a contact‐dependent mechanism to directly and actively influence the immune response.

Catheter‐associated urinary tract infections account for up to 12% of hospital‐acquired infections and are associated with a mortality rate of approximately 2.3%, or up to 10% when bacteraemia is an outcome [87]. This significant clinical burden has driven intensive research efforts toward antibacterial and/or antifouling surface modifications of catheter properties [88]. In one approach, a group fabricated a bactericidal coating consisting of a polystyrene matrix that encapsulated silver nanoparticles that were capped with ricinoleic acid [89]. To further enhance the antibacterial activity of the coating, the group loaded tetracycline into the polystyrene matrix. The group reported potent antibacterial properties, with a bactericidal activity above 90%, which was attributed to a burst release of tetracycline and a gradual release of silver ions. This activity profile would be attractive during the initial hours of catheterization; however, the effect may be limited due to the exhaustion of the loaded therapeutics. In a separate approach, another group developed a dual antimicrobial and antifouling coating by co‐depositing a quaternary ammonium polyethyleneimine (QPEI) with dopamine and poly(carboxylbetaine‐co‐dopamine methacrylamide) (pCBDA) [90], which they referred to as a ‘QCB’ coating. Within the architecture of the QCB coating, the QPEI component served as a contact‐dependent bactericidal functionality, while the pCBDA component provided antifouling properties. The authors showed that the QCB coating could significantly reduce bacterial adhesion and effectively kill many of the remaining bacteria that successfully attached. In this regard, the antimicrobial activity of the QCB coating can be considered more attractive than that of the nanoparticle/tetracycline‐embedded polystyrene coating, principally because its contact‐dependent nature may prolong the activity of the effect. This rationale was also applied to our study, which relies on contact between the functional moieties on the p(PEO) coating and the adherent bacterial cell.

In addition to minimizing bacterial adhesion, the QCB surface was shown to reduce immune cell infiltration into the bladders of rabbits in vivo [90]. This passive anti‐inflammatory effect was likely driven by the antifouling characteristic of the QCB coating, which reduces protein adsorption, thereby delaying the critical first step in the immune recognition of biomaterials [91]. Similarly, related studies have also attributed reduced immune activation to the suppression of protein adsorption [92]. However, while a reduction in protein adsorption may passively delay the immune response to the foreign material, it would not actively alleviate the mechanically triggered inflammatory response that frequently occurs during bladder catheterization [93]. This sterile inflammatory response is reported to occur following abrasion of the epithelium during catheter insertion. A more suitable material coating should possess the functionality to actively resolve the inflammatory response by suppressing the expression of pro‐inflammatory cytokines and shifting macrophage polarization to the M2 state. Toward this, our p(PEO) coating has demonstrated the promising ability to actively modulate the immune system. This effect is potentially driven via interactions between eucalyptol‐derived moieties in the polymer network and PTK2B bound to the immune cell membrane [85].

The present study yielded favourable results that collectively promote p(PEO) as a promising bioactive coating for reducing complications associated with catheter applications. The multifunctionality of the coating enables it to simultaneously resolve a sterile inflammatory response, eliminate adherent bacteria, and promote the activity of prophylactic antibiotics. This represents multiple advancements in the field. First, the contact‐dependent immunomodulatory attribute is an attractive advantage over other coatings that either passively reduce protein adhesion [90] or rely on diffusing anti‐inflammatory drugs [94]. This property ideally grants the p(PEO) coating the ability to resolve local abrasion‐related inflammation following catheter insertion, while not releasing therapeutics systemically. Another attractive advancement of the p(PEO) coating is its demonstrated ability to sensitize adherent pathogens to conventional antibiotics, driven by the permeabilization of sub‐lethally injured cells. This is a desirable trait for implantable biomaterials in general, due to the common practice of administering prophylactic antibiotic treatments upon implantation. In addition, the coating fabrication described in this report drives an advancement toward greener biomedical technologies. This is supported by the renewably sourced peppermint essential oil precursor and coating deposition by solvent‐free plasma‐polymerization. Lastly, while we have designed our research and interpreted our findings in the context of catheter‐associated urinary tract infection, the substrate‐independent nature of plasma‐polymerization enables the approach to be adapted to other biomaterials and implanted devices. To validate this, we deposited the coating onto Ti6Al4V (Figure S9), which is commonly used in orthopaedic implant applications. To evaluate whether the coating retains a similar performance on the Ti6Al4V substrates, we measured water contact angle, antibacterial activity against Gram‐negatives, and radical scavenging activity. Analyses of these properties were consistent with the results presented for the silicon wafers and silicon tubing, indicating that the properties of the bulk material have little effect on the coating deposition and performance. Our coating was durable in aqueous conditions and non‐haemolytic, so that it may also be suitably applied to central venous catheter applications. Future research in this field will benefit from exploring plasma deposition of varying essential oils for application across multiple implanted devices.

3. Conclusion

This study examines the application of atmospheric plasma deposition of peppermint essential oil, p(PEO), as a biocompatible and multifunctional coating for medical devices. We optimized plasma process parameters to create a durable substrate‐independent coating with antioxidant, antibacterial, and immunomodulatory properties. This coating was shown to retain diverse precursor‐derived functional groups that drive bioactive properties. We evaluated its performance within a clinical context of bladder catheterisation as a pilot application, as this is an area of medicine that is strongly burdened by oxidative stress, inflammation, and infection.

The p(PEO) coating was shown to scavenge up to 90% of reactive oxygen species, underscoring its considerable antioxidant activity. In parallel, the coating was shown to have a specific antibacterial activity against Gram‐negative pathogens, including P. aeruginosa and two E. coli isolates (including a standard laboratory strain and a clinically obtained uropathogenic isolate). We determined that the antibacterial activity of the p(PEO) coating is driven by a contact‐dependent disruption of the Gram‐negative cell membrane, causing perforation and a loss of membrane polarization. To build on this, we investigated whether the coating activity could be coupled with antibiotic therapy to improve antibacterial outcomes further. To do so, we utilized two antibiotics, colistin and levofloxacin, which are frequently used to treat and prevent urinary tract infections. Our results showed that the bacteria cultured on the multifunctional coating were sensitized to both antibiotics – an effect which we attribute to the high rate of bacterial cell permeabilization and the downstream stresses that subsequently emerge. The immunomodulatory properties of the p(PEO) coatings were reflected in their ability to reduce the expression of IL‐6 and TNF‐α and increase the expression of IL‐4. In addition, we utilized fluorescence‐based analyses to observe a shift in macrophage polarization to M2 following incubation on p(PEO), further supporting the resolution of inflammation. Overall, these in vitro results are highly encouraging because they indicate that urinary catheters coated with p(PEO) could be deployed to support patient recovery in multiple ways. First, the p(PEO) catheter may mitigate the sterile inflammatory response commonly associated with bladder catheterization. In addition, the coated device could act as a strong defence against catheter‐associated urinary tract infections through its inherent antibacterial activity and its ability to potentiate prophylactic antibiotic treatment. Lastly, from an engineering perspective, the findings of this research underscore the untapped potential of employing atmospheric plasma deposition of plant‐based essential oils to advance biomedical engineering within a context of sustainability and a low‐carbon footprint economy.

4. Experimental Section

4.1. Materials

Peppermint oil (Essential Health, ALDI, Australia, AUST L 319481), DPPH, and ABTS (Sigma–Aldrich). LIVE/DEAD Bacterial Viability Kit (BacLight, Invitrogen), H2DCFDA (Thermo Fisher), 4% glutaraldehyde (Sigma–Aldrich Australia), paraformaldehyde (ThermoFisher), DAPI and phalloidin, Dulbecco's Modified Eagle's Medium (DMEM), fetal bovine serum (FBS), penicillin–streptomycin solution, trypan blue, trypsin,(Sigma–Aldrich), Ultrahigh purity water gained from a MilliQsystem (Millipore Milli‐Q Academic, USA), phosphate buffered saline tablet (Sigma–Aldrich), MTT assay kit (Sigma, Darmstadt, Germany). IL‐6 and IL‐4 ELISA kits (Invitrogen, USA), anti‐CD80‐Alexa Fluor 488 (Abcam, UK), anti‐CD206‐FITC (Abcam, UK), anti‐hu CD206‐FITC (Invitrogen, USA). Silicone catheter (Livingstone, Australia). ROS‐Glo H2O2 Assay kit (Sigma–Aldrich, USA).

4.2. Atmospheric Plasma Deposition of p(PEO)

The configuration of the integrated atmospheric plasma systems for the deposition of peppermint essential oil is illustrated in Figure 1A and Figure S10. Two designs of in‐house plasma reactors were utilized to accommodate samples of varying sizes and geometries.

To apply a coating on silicon wafers, silicone tubing, and titanium coupons, we utilized the arrangement illustrated in Figure S10A. This design utilized an AC power supply with an electronic transformer (NP‐10000‐30, NeonPro Co, USA) to energize the chamber coating system. The power supply sustained a constant frequency of 24 kHz, with the voltage established at 5 kV. A chamber housing two tungsten‐steel rods (150.0 mm × 2.4 mm) was placed within a quartz tube (200 mm in length, with an inner diameter of 20.0 mm and an outer diameter of 25.0 mm), maintaining a separation of 13 mm between the rods. The chamber was linked to the high‐voltage power source, and a circular grounded electrode was situated 15 mm from the nozzle's open end. This apparatus facilitates the generation of plasma by dielectric barrier discharge (DBD). A 10 kΩ resistor was fitted to avoid plasma flashback into the chamber. The substrates designated for coating were positioned within the chamber, with adjustable argon gas provided and linked to the peppermint oil glass container, facilitating the amalgamation of the argon carrier gas with the oil vapor, which was subsequently introduced into the plasma chamber. The plasma was immediately ignited from the electrodes positioned 20 mm above the samples for durations of 5 or 10 min.

To facilitate coating of the well plates, the plasma nozzle was utilized to deposit the coating onto the surface of the well, illustrated in Figure S10B. A solitary tungsten‐steel rod (150.0 mm × 2.4 mm) served as the high‐voltage electrode in this design. It was positioned within a glass tube (200 mm in length, with an inner diameter of 8.0 mm and an exterior diameter of 10.0 mm) that functions as a gas flow conduit. The outermost glass tube, measuring 17 mm in length with an inner diameter of 20.0 mm and an outside diameter of 22.0 mm, was inserted to form the dielectric layer, while a circular grounded electrode was placed 15 mm from the open end of the tube. The inner tube was secured with a 3D printed Thermoplastic Polyurethane (TPU) insulator with an inner diameter of 10.0 mm and an outside diameter of 20.0 mm. A TPU cap was used to link components together. To produce the plasma, the sample power supply and resistor were connected as outlined in the initial configuration. The working distance remained unchanged. In order to validate that the two plasma system configurations would produce equivalent coatings, we measured the coating thickness and found no significant difference between plasma configurations.

4.3. Optical Emission Spectroscopy

The optical sensor head was positioned 200 mm from the plasma jet. A PLASUS EMICON MC spectroscopic plasma monitor and process control system (PLASUS, DE) with EMICON MC version 4.6 software recorded the optical emission spectroscopy of either pure Argon gas or a mixture of Argon gas and essential oil plasma.

4.4. Coating Thickness

The coating thickness was measured by a spectroscopic ellipsometer (SENresearch 4.0, SENTECH Instruments, GmbH, Berlin, Germany). Measurements were performed with a variable‐angle spectroscopic ellipsometer imaging system. The change in the polarisation state of light upon reflection from the film surface was processed using SpectraRay/4 comprehensive ellipsometry software (SENresearch 4.0) to evaluate the coating's thickness. Each coating condition was replicated four times for the thickness measurement.

4.5. Stability Test

The thickness of the coatings was measured before and after being submerged in phosphate‐buffered saline (PBS; isotonic solution) and Milli‐Q water for 24 h at room temperature. The reduction percentage was calculated using the following equation:

$$\begin{array}{c}\hfill \mathrm{Thickness}\mathrm{reduction}\mathrm{percentage}\left(\%\right)=\\ \hfill \frac{thickness\left(before\right)-thickness\left(after\right)}{thickness\left(before\right)}\times 100\end{array}$$

4.6. Wettability

The wettability of the plasma‐modified surfaces was measured using a sessile water drop contact angle goniometer (RD‐SDM02, RD Support, Scotland, UK). 5 µL of Milli‐Q water was dropped onto the plasma‐modified substrates, and an image was immediately captured upon contact. ImageJ v1.53 analysis software (NIH, MD, USA) was used to measure the tangent of the drop at its three‐phase contact point (intersection between solid, liquid, and gas).

4.7. Zeta (ζ) Potential

A ZPA 2.0 (Dataphysics, Germany) was used to measure the ζ‐potential of the plasma‐deposited substrates over a pH range of 3–9. An oscillating flow of KCl(aq) (10⁻³ m) was applied in a narrow gap of approximately 130 µm between two p(PEO)‐coated substrates with a defined area of 10 × 20 mm. An automated dosing unit adjusted the pH between each reading using stock solutions of 1 m HCl(aq) and 1 m KOH (aq).

4.8. Fourier‑Transform Infrared (FTIR) Spectroscopy

FTIR measurement was performed to determine the chemical properties of the plasma‐deposited polymers. Briefly, the coated silicon wafer (1 × 1 cm) was coated with p(PEO) and kept stable in an air‐prohibited vacuum seal bag for 24 h before the measurement. The samples were later loaded into the ATR‐FTIR holder of a PerkinElmer Spectrum 3 FTIR spectrometer (PerkinElmer, MA, USA). The spectra were collected with 128 scans ranging from 500 to 4000 wavenumber (cm⁻¹).

4.9. X‑Ray Photoelectron Spectroscopy (XPS)

The surface chemical composition of p(PEO)‐coated silicon wafers was analyzed using a Kratos AXISUltra DLD spectrometer. X‐ray photoelectron spectroscopy (XPS) spectra were recorded with a monochromatic AlKα (hν = 1486.7 eV) radiation source conducted at an electric current of 15 mA and a voltage of 15 KeV. Survey spectra were recorded over a range of 0–1100 eV at a pass energy of 160 eV, and high‐resolution spectra were performed at 20 eV. Casa XPS software was used for data analysis. All binding energies were referenced to the aliphatic carbon C1s peak at 285.0 eV. Curve fitting of core‐level envelopes was performed with the minimum number of Gaussian–Lorentzian components, generally at 30% Lorentzian and 70% Gaussian functions.

4.10. Atomic Force Microscopy

To examine the coatings' surface topography, the coated silicon wafers (5 and 10 min) were measured by Bruker Dimension Icon AFM in ScanAsyst‐air mode. Then, the surface topography images were constructed and analyzed by Gwyddion software (2.68, Czech Metrology Institute).

4.11. Bacterial Culture and Testing Conditions

The non‐coated substrates (silicon wafer or silicone catheter tube) and the coated samples were tested for antibacterial activity against four bacterial strains: S. aureus ATCC25923, S. epidermidis ATCC 14990, E. coli ATCC11303 (B), and P. aeruginosa ATCC15692. In addition, the p(PEO) modified catheter tubes were evaluated for their antibacterial activity against a clinical isolate of uropathogenic E. coli, supplied by SA Pathology, Flinders Medical Centre. Bacterial strains were recovered from glycerol stocks stored at −80°C and streaked for purity on tryptone soy agar (TSA). One isolated colony of each species was transferred aseptically from TSA to 5 mL of tryptone soy broth (TSB) and cultured at 37°C until the late log phase (≈18 h). The coated substrates were aseptically placed in sterile 24‐well plates and immersed in 0.5 mL of the 10⁵ CFU mL⁻¹ bacterial suspension and then incubated for 6 or 24 h.

4.12. Colony Enumeration

Following bacterial inoculation and incubation, samples were vortexed in PBS for 15 s, followed by sonication for 5 min before being vortexed for another 15 s, then serially diluted 10‐fold. In triplicate, serially diluted samples (10 µL) were dropped onto TSA plates and incubated for 18 h at 37°C. Standard plate counts evaluated viability, and the quantity of colony‐forming units (CFU) per sample was calculated using the number of colonies counted, the aliquot volume, and the dilution factor.

4.13. Live/Dead Staining

Overnight bacterial cultures were diluted with TSB to obtain 1 × 10⁶ colony‐forming units/mL (CFU/mL). The diluted bacterial suspensions were seeded on the untreated silicon wafers and p(PEO) coated silicon wafers. The samples were then incubated for 6 h or 24 h. They were washed two times with PBS and stained with the LIVE/DEAD BacLight viability kit (Invitrogen, MA, USA), following the manufacturer's instructions, for 10 min at room temperature under dark conditions and immediately imaged in triplicate with a ZEISS LSM 880 confocal laser scanning microscope (Zeiss, Germany), using excitation/emission settings of 480/500 nm for SYTO9 and 490/635 nm for PI.

4.14. Scanning Electron Microscopy

Morphological alterations in bacteria were observed utilising FEI Inspect F50 field emission scanning electron microscopy (SEM). The sample preparation for SEM included glutaraldehyde fixation in 0.1 m sodium cacodylate buffer (pH 7.4) for 45 min. The samples were further dehydrated utilising a graded ethanol series (70%, 80%, 90%, and 100%) for 10 min. The specimens were dehydrated using nitrogen gas and subsequently sputter‐coated with platinum utilising an ion sputter coater, TB‐SPUTTER (Quorum Technologies, UK).

4.15. Cell Culture

Human monocytic cells (THP‐1) were grown in RPMI 1640 (Thermo Fisher Scientific, MA, USA), with supplements for growth, 10% v/v fetal bovine serum (FBS; Gibco, MT, USA), and contamination prevention, and 1% v/v penicillin/streptomycin. Cells were cultured at 37°C in a 5% CO₂ incubator until the desired cell number of 10⁶ cells was reached. For cell adhesion, differentiated THP‐1 cells were used to investigate the impact of surfaces on immune cells. The differentiation of THP‐1 cells was induced by PMA 100 ng/mL for 48 h, followed by a 24 h rest period with PMA‐free DMEM (Thermo Fisher Scientific, MA, USA). The cells were detached by 0.25% trypsin and neutralized by DMEM. The suspension was centrifuged to remove the remaining trypsin and resuspended in DMEM. Single‐cell suspension was counted before being seeded on surfaces at a ratio of 0.8 × 10⁵ viable cells per surface. Surfaces were incubated at 37°C in a 5% CO₂ atmosphere for 24 h (for MTT assay, cell imaging, DCF assay, ELISA, and RNA expression), 48 and 72 h for MTT assay.

The HaCaT cells (human epidermal keratinocyte line, Cell Line Services, Eppelheim, Germany) were grown in Dulbecco's Modified Eagle's Medium (DMEM) supplemented with 10% heat‐inactivated fetal bovine serum (FBS, Gibco, Australia) and 1% streptomycin/penicillin (Gibco‐BRL, Australia). Upon achieving around 80%–85% confluence (around 10⁶ cells), the cells were collected utilizing 0.25% Trypsin‐EDTA (1X) (Gibco‐BRL, AU). A volume of 200 µL of HaCaT cells, at a density of 1 × 10⁵ cells mL⁻¹, was inoculated onto the substrates. Substrates were incubated at 37°C in a 5% CO₂ atmosphere for 24 h for the MTT assay and cell imaging. To capture the cell images, the substrates were rinsed with PBS to remove unbound cells, then were stained with DAPI (excitation at 358nm/ emission at 461nm) and Phalloidin‐FITC (496nm/ 561nm) for a duration of 20 min for each stain. Finally, the samples were examined with a ZEISS LSM 880 confocal laser scanning microscope (Zeiss, Germany).

4.16. MTT Assay

According to the manufacturer's instructions, cell viability was assessed utilizing the MTT assay kit (Sigma, Darmstadt, Germany). MTT (3‐(4,5‐dimethylthiazol‐2‐yl)‐2,5‐diphenyl tetrazolium bromide; Sigma) was dissolved in PBS at a concentration of 5 mg/mL, and passed through a 0.22 µm filter to sterilize and remove minor insoluble particles. Stock MTT solution (10 µL per 100 µL media) was administered to all wells of an assay, followed by incubation of the plates at 37°C for 4 h. DMSO (ChemSupply, Australia) was added to each well to dissolve the dark blue crystals and was thoroughly mixed. The plates were analyzed using a SYNERGY‐HTX multi‐well plate reader (Bio‐Tek, Vermont, USA) at a test wavelength of 570 nm following a brief incubation at room temperature to confirm complete dissolution of all crystals. The percentage of cell viability was determined by normalizing the results against control cells cultured in medium only. Wells containing only culture media represent 100% cell viability.

4.17. DCF Assay

The DCF (dichlorofluorescein) staining assay is a technique employed to quantify reactive oxygen species (ROS) in cells. This process utilizes a non‐fluorescent molecule, DCFH‐DA (dichlorodihydrofluorescein diacetate), which permeates cells and is transformed into a fluorescent variant, DCF, through oxidation by ROS. The fluorescence intensity is subsequently associated with the quantity of ROS present. THP‐1 and HaCaT cells, individually, were seeded onto the testing substrates at a density of 1 × 10⁵ cells and cultured at 37°C with 5% CO₂ overnight. The samples were then rinsed three times with PBS, followed by adding DCFA solution (20 µm DCFDA in PBS solution) and then incubated at 37°C in the dark for 40 min. Before imaging, the supernatant was discarded, and the cells were rinsed with PBS. Subsequently, DAPI was incorporated into the samples, and the substrates were imaged using a ZEISS LSM 880 confocal laser scanning microscope. DCF fluorescence was measured using Ex/Em = 485/535 nm, while DAPI fluorescence was measured using Ex/Em = 485/535 nm.

4.18. ELISA Assays for Pro‐Inflammatory and Anti‐Inflammatory Cytokines

IL‐6 and IL‐4 ELISA kits (Invitrogen, USA) were used based on the manufacturer's instructions. After three washes with washing solution, samples or standards are introduced, followed by a 2 h incubation at ambient temperature. After further washing, a streptavidin‐conjugated horseradish peroxidase is introduced and incubated for 1 h at room temperature, facilitating signal amplification. The wells are rewashed, subsequently incorporating a chromogenic TMB substrate. Following a 30 min incubation in darkness, the enzymatic process is terminated, and the absorbance at 450 nm is assessed using a spectrophotometer to quantify the target cytokines.

4.19. RNA Extraction

Substrate‐attached cells were gently rinsed with PBS to eliminate non‐adherent cells, then detached using ultrasonication for 2 min, followed by 30 s of vortexing. The cells were pelleted via centrifugation and resuspended in an RNA extraction buffer provided by the RiboPure Bacteria RNA kit (Invitrogen, Massachusetts, USA). RNA extraction was performed following the manufacturer's guidelines. The quantity of RNA was measured, and its quality was assessed using a Nanodrop 2000c spectrophotometer (ThermoFisher, Massachusetts, USA).

4.20. RT‐PCR

PCR master mixes were created using the SuperScript III Platinum One‐Step qRT‐PCR Kit (Invitrogen, MA, USA), with primers included at a concentration of 10 µM. The housekeeping gene is GADPH (Genbank ID: NM_001357943). The primer sequences are provided in Table S2. Each reaction tube received 1 ng of template RNA in 1 µL aliquots, while no‐template controls were given 1 µL of RNAse‐free H₂O instead. Reverse transcription and amplification occurred in a single step on a Rotor‐Gene Q Thermocycler (version 2.1.0, QIAGEN, Hilden, Germany) using the following protocol: 3 min at 50°C, 5 min at 95°C, and 40 cycles of 95°C for 15 s and 60°C for 30 s. The signal was collected at 60°C, and a melt curve was produced in 1°C increments from 72°C to 95°C.

4.21. Pro‐Inflammatory Marker (CD80) and Anti‐Inflammatory Marker (CD206) Staining

To determine the ratio of pro‐inflammatory to anti‐inflammatory markers present on macrophages, we looked for the ratio of CD206 signal/ CD80 signal, since CD206 is the marker of anti‐inflammatory M2 phenotype macrophages [95, 96] while CD80 is the marker for pro‐inflammatory M1 phenotype macrophages [82, 97]. To begin the experiment, 2 × 10⁵ macrophages were seeded onto the samples with or without adding LPS (lipopolysaccharide) and cultured overnight at 37°C with 5% CO₂ before the experiment. Subsequently, 50 µL of anti‐CD80‐Alexa Fluor 488 (Abcam, UK) was applied to the surfaces and incubated at 37°C for 30 min. The mixture was subsequently rinsed twice with PBS. Subsequently, 50 µL of anti‐CD206‐FITC (Abcam, UK) was added to the samples and incubated at 37°C for 30 min. Subsequently, all examined samples were rinsed twice with PBS and imaged using a ZEISS LSM 880 confocal laser scanning microscope (Zeiss, Germany).

4.22. Flow Cytometry of Anti‐Inflammatory Marker (CD206)

We employed CD206 biomarker labelling to quantify the polarized M2 phenotype macrophages [95, 96]. In summary, 2 × 10⁵ THP‐1 macrophages were inoculated onto the experimental surfaces with or without LPS and cultured overnight at 37°C in a 5% CO₂ atmosphere. The cell suspensions were subsequently gathered. The samples were subsequently stained with anti‐hu CD206‐FITC (Invitrogen, USA) at 5µL of dye per 1 mL of cell suspension. The samples were subsequently analyzed using the flow cytometer (Cytoflex S, Beckman Coulter, USA) to capture the FITC signal throughout a sample size of 10 000 events.

4.23. Antioxidant Activity Determinations

The stock solutions for the 2,2'‐azino‐bis(3‐ethylbenzothiazoline‐6‐sulfonic acid) (ABTS) test comprised a 7.4 mm ABTSd+ solution and a 2.6 mm potassium persulfate solution. The operational solution was subsequently formulated by combining the two stock solutions in equal volumes and permitting them to react for 12 h at ambient temperature in the absence of light. The solution was further diluted by combining 1 mL of ABTS^*+ solution with 60 mL of methanol to achieve an absorbance of 1.170 ± 0.02 units at 734 nm, as measured by the spectrophotometer. A new ABTS^*+ solution was produced for each test. The testing substrates were permitted to interact with 2850 mL of the ABTS^*+ solution for 30 min or 2 h in a dark environment. The absorbance was measured at 734 nm with the spectrophotometer. 2850 mL of the stock ABTS was used as the control and kept in the same experimental condition for 30 min or 2 h [98]. The following formula calculated the scavenging activity:

$$\begin{array}{c}\hfill \text{Scavenging activity}\left(\%\right)=100\times \left[\left({\mathrm{A}}_{734}\text{of the}\mathrm{control}\right.\right.\\ \hfill -\left.\left.{\mathrm{A}}_{734}\text{of the sample}\right)/\left({\mathrm{A}}_{734}\text{of the control}\right)\right]\end{array}$$

A stock solution of 2,2‐diphenyl‐1‐picrylhydrazyl (DPPH) was produced by dissolving 24 mg of DPPH in 100 mL of methanol and subsequently kept at −20°C until required. A working solution was prepared by combining 10 mL of stock solution with 45 mL of methanol, resulting in an absorbance of 1.1 ± 0.02 units at 515 nm, as measured by the spectrophotometer. The samples were permitted to react with 2850 mL of the DPPH solution. 2850 mL of the stock DPPH solution was utilised as the control and maintained under identical experimental conditions for 3 or 6 h in the dark, after which the absorbance was measured at 515 nm [98]. The scavenging activity was determined using the subsequent formula:

$$\begin{array}{c}\hfill \mathrm{Scavenging}\mathrm{activity}\left(\%\right)=100\times \left[\left({\mathrm{A}}_{515}\mathrm{of}\mathrm{the}\mathrm{control}\right.\right.\\ \hfill -\left.\left.{\mathrm{A}}_{515}\mathrm{of}\mathrm{the}\mathrm{sample}\right)/\left({\mathrm{A}}_{515}\mathrm{of}\mathrm{the}\mathrm{control}\right)\right]\end{array}$$

4.24. Statistical Analysis

All experiments were performed using an independent‐groups design in which each treatment condition represented a separate experimental group. A one‐way ANOVA was used to compare means across three or more groups, followed by Bonferroni‐corrected post hoc tests to account for multiple pairwise comparisons. For cytocompatibility studies, raw viability measurements of each treatment group were normalized to the measurements of the untreated cells incubated in tissue culture plates. Statistical significance was defined as p < 0.05. All analyses were performed using GraphPad Prism (v10.4.2, GraphPad Software Inc., California, USA). For each condition, n = 3 independent biological replicates were analysed. Data are presented as mean ± standard deviation (SD).

Funding

NHMRC for Fellowship GNT1194466; ARC for grants DP220103543 and DP250101028. ARC for grant FT240100067. Flinders Foundation for Health Seed Grant.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File: smll72405‐sup‐0001‐SuppMat.docx.

Data Availability Statement

Data will be made available upon reasonable request to the corresponding authors.