Transparent polycarbonate coated with CeO₂ nanozymes repel Pseudomonas aeruginosa PA14 biofilms

Abstract

Highly transparent CeO₂/polycarbonate surfaces were fabricated that prevent adhesion, proliferation, and the spread of bacteria. CeO₂ nanoparticles with diameters of 10–15 nm and lengths of 100–200 nm for this application were prepared by oxidizing aqueous dispersions of Ce(OH)₃ with H₂O₂ in the presence of nitrilotriacetic acid (NTA) as the capping agent. The surface-functionalized water-dispersible CeO₂ nanorods showed high catalytic activity in the halogenation reactions, which makes them highly efficient functional mimics of haloperoxidases. These enzymes are used in nature to prevent the formation of biofilms through the halogenation of signaling compounds that interfere with bacterial cell–cell communication (“quorum sensing”). Bacteria-repellent CeO₂/polycarbonate plates were prepared by dip-coating plasma-treated polycarbonate plates in aqueous CeO₂ particle dispersions. The quasi-enzymatic activity of the CeO₂ coating was demonstrated using phenol red enzyme assays. The monolayer coating of CeO₂ nanorods (1.6 μg cm⁻²) and the bacteria repellent properties were demonstrated by atomic force microscopy, biofilm assays, and fluorescence measurements. The engineered polymer surfaces have the ability to repel biofilms as green antimicrobials on plastics, where H₂O₂ is present in humid environments such as automotive parts, greenhouses, or plastic containers for rainwater.

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Introduction

Polycarbonate (PC) is an engineering plastic with a wide range of applications that is used as a substitute for glass¹ for large-area coverings and safety glazing, e.g., in the automotive industry, in greenhouses, or for outdoor covers, as it is much softer and easily moldable to meet modern design and production requirements. However, low hardness, scratch resistance, and low resistance to photo- or biocorrosion limit the use of PC for many industrial applications. It is therefore of interest to modify or improve the surface of the polymer without changing its bulk properties. PC is very transparent to visible light and has better light transmission properties compared to different types of silica glass. However, PC unfortunately has a low glass transition temperature of ∼150 °C and gradually softens above this temperature, and begins to flow above ∼300 °C. To allow PC to be used in applications exposed to weathering or UV radiation, a special surface treatment is required. The surface properties of plastic surfaces can be specifically modified by coating. There are a variety of strategies to functionalize polymer surfaces for biomedical applications by plasma treatment, superhydrophobic surfaces, quaternary ammonium compounds, chitosan-containing surfaces, or embedded nanoparticles (NPs).^2–7

Coating polymer surfaces with metal oxide nanoparticles allows potentially interesting commercial applications such as self-cleaning or antibacterial properties.⁸ Polymer surfaces coated with TiO₂ nanoparticles are more scratch-resistant and have antibacterial properties due to the photocatalytic properties of TiO₂ but a recent classification of nano-TiO₂ as a “suspected carcinogen” by the EU may lead to restrictions or even a ban on their chemical use in consumer products.⁹ Coating with ZnO/TiO₂ NPs may lead to self-cleaning, while coating with nano-Ag or carbon nanotubes may confer antibacterial properties.¹⁰ A common problem with silver nanoparticles or antibiotics is the decrease in the antibacterial activity due to the leaching of the active compound with time, leading to materials with a limited life span and metal (Cu, Ag) nanoparticles that are harmful to the environment.¹¹ Tailored surfaces to which bacteria and cells cannot adhere represent an alternative approach to combat bioadhesion.¹² Surfaces coated with poly(ethylene oxide) or polymers with zwitterionic groups are known to suppress nonspecific protein adsorption and bacterial adhesion.^13,14 Superhydrophobic antibacterial coatings with polycarbonate have been proposed for catheters to prevent infections.¹⁵

Antibiotics are the most important tools in the fight against bacteria. However, bacteria are adaptable and can build up resistance to antibiotics.¹⁶ For this reason, research is constantly being conducted to find other ways of combating them. One of these antibiotic-resistant bacteria is the Gram-negative bacterium Pseudomonas aeruginosa (P. Aeruginasa). It is among the most common causes of respiratory pneumonia, wound infections, and urinary tract infections worldwide.¹⁷

Our approach to fabricating biofilm-inhibiting surfaces of such bactaria is non-toxic and inspired by nature: using oxidizing halogenating enzymes (haloperoxidases, HPOs), seaweeds catalyze the oxidation of halides (Cl⁻ and Br⁻) with hydrogen peroxide (H₂O₂) to form hypohalous acids (HOCl and HOBr).^18–20 In the presence of ubiquitous Br⁻ ions and H₂O₂, generated in sunlight, CeO₂ is—by virtue of its catalytic activity—a functional substitute for haloperoxidases, which can also brominate AHL signal molecules so that communication between the bacteria is disrupted (quorum quenching) and biofilm formation is inhibited in a non-biocidal way.²¹ Metal complexes,^22–24 V₂O₅,²⁵ and CuO²⁶ or ceria^21,27 nanoparticles have been reported to act as “nanozymes” (inorganic nanoparticle-based artificial enzymes) by mimicking the HPO activity.^28,29 Ceria nanoparticles are readily and inexpensively available, non-toxic,³⁰ highly insoluble (pK_L = 60),³¹ stable over a wide pH range,³² and therefore, environmentally safe. Thus, they are ideally suited for real-world applications.^22,33–35

The important problems in the coating of polymer surfaces are the low temperature stability and the low surface energy of the polymer. These lead to poor surface wetting, which results in weak adhesion.³⁶ Polymer surfaces can be coated with metal oxides in various ways to achieve potentially interesting commercial applications such as self-cleaning or antibacterial properties.³⁷ Different low-temperature techniques such as pulsed laser deposition (PLD),³⁸ magnetron sputtering,³⁹ spray pyrolysis,⁴⁰ and metal organic chemical vapor deposition (MOCVD)⁴¹ are available to prepare surface-active CeO₂ films at low temperature. However, most physical vapor deposition processes have problems in depositing large area films, require complex optimization of deposition conditions, and the substrates are silicon or transition metals. Wet chemical deposition is suitable for producing thin layers on large surfaces but drying metal oxide films on heat-sensitive polymer substrates is problematic⁴² as it leads to thin films with low surface area and low catalytic activity. A quick and simple method to increase the surface energy and adhesion is plasma treatment with oxygen (O₂) or argon (Ar).^33,41 As a result, oxygen-containing groups such as hydroxyl, carbonyl, and carboxyl groups are incorporated on the surface, thus increasing the polarity.⁴²

Last but not least, CeO₂ nanoparticles for surface coatings must be available in sufficiently large quantities and be well dispersible to achieve homogeneous coatings as agglomerates otherwise impair the optical appearance and deteriorate the surface roughness of the material. Since nanoparticles are only reactive when exposed at the surface, coating approaches are much more material efficient and effective than embedding particles in a bulk polymer.²¹ We present (i) a large scale synthesis of CeO₂ nanorods along with (ii) a polycarbonate coating approach that allows easily scalable process parameters and thus potential mass production. (iii) We demonstrate by scanning electron microscopy and scanning probe techniques the bacterial repellancy of the film and (iv) show the quorum quenching mechanism through the production of the virulence factor pyocyanin in response to the haloperoxidase-like activity of the CeO₂ coating. The proven non-toxicity of the coating makes polycarbonate surfaces promising for resisting biofilm formation in daily use where H₂O₂ is present in humid environments such as greenhouses or plastic containers for rainwater (H₂O₂ production in ground water,⁴³ the atmosphere,⁴⁴ or in natural water exposed to sunlight).⁴⁵

Results and discussion

Particle preparation and analysis

A freshly prepared cerium(III) hydroxide dispersion was used for the synthesis of ceria nanoparticles in aqueous media with hydrogen peroxide as the oxidant and nitrilotriacetic acid (NTA) as the capping agent. The capping agent ensured the formation of a stable suspension without inhibiting the haloperoxidase activity of the CeO₂ nanoparticles by forming surface complexes. In addition, the capping agent can be used to produce a stable suspension that is used for polycarbonate coating. The reaction can be easily scaled up to the 100 g scale in batch operation for coating large surface areas.

Fig. 1A displays the TEM micrograph of a sample obtained after centrifugation from an aqueous nanoparticle dispersion. The reaction yielded nanorods with a length of 100–200 nm and a width of 10–15 nm, suitable for the transparent coating of surfaces. The X-ray powder diffraction pattern confirmed the formation of single phase CeO₂ (Fig. 1C) in the bulk sample. The nanoparticles were carefully washed three times by centrifugation at 9000 rpm for 30 min. We avoided irreversible agglomeration by preventing complete solvent evaporation. NTA is well-suited as a ligand for CeO₂ because the carboxylate groups are hard donors for ionic bonding to Ce⁴⁺ ions. NTA is thought to form a complex with cerium surface atoms, in which the three carboxylate groups are bound by ionic interactions and the N atom of the NTA ligand by dative interactions.⁴⁶ Light scattering experiments revealed an average hydrodynamic radius of 115 ± 79 nm for CeO₂ nanoparticles with NTA surface ligands.

Fig. 1 (A, B) TEM image of the CeO₂ nanorods and (C) X-ray powder diffraction pattern of a CeO₂ sample. Red ticks indicate the calculated reflection positions for CeO₂ based on the structural data from the JCPDS data bank (JCPDS# 00-034-0394).

Haloperoxidase activity

The haloperoxidase-like activity of CeO₂ nanorods was demonstrated with a phenol red bromination assay. In this assay, a phenolsulfonphthalein indicator (phenol red, PR) is brominated in aqueous solution in the presence of Br⁻ and H₂O₂ to bromophenol blue (3′,3′′,5′,5′′-tetrabromophenolsulfonphthalein, Br₄PR). The reaction can be monitored spectrophotometrically because the educt and product have different distinct absorption maxima at λ_max,PR = 430 nm, and λ_max,Br₄PR = 590 nm. Michaelis–Menten enzyme kinetics were assumed because the haloperoxidase-like activity was dependent on the H₂O₂ concentration. The kinetics were measured for different concentrations of H₂O₂ and fitted with the Hill equation (eqn (1)) (Fig. 2).⁴⁷

Fig. 2 Haloperoxidase-like activity of CeO₂ nanoparticles determined with a phenol red bromination assay. (A) Time-dependent UV-vis spectra indicate the change in the oxidative bromination from phenol red (PR) to bromophenol blue (Br₄PR). (B) Shows the Michaelis–Menten kinetics for the substrate H₂O₂ using the CeO₂ enzyme mimic.

The Michaelis constant K_m is a measure of the affinity of the enzyme, in this case CeO₂, for the H₂O₂ substrate with the concentration [S]₀. The value K_m has a reciprocal relationship with the affinity, i.e., the higher the constant, the weaker the binding strength between the enzyme and the substrate. For CeO₂, the constant is 0.85 mM, which indicated a high affinity for H₂O₂. The maximum rate of the reaction is described by the v_max value, which is 0.021 μM min⁻¹ for the CeO₂ nanorods.

The bromination rate depends on (i) the surface charge, (ii) BET surface area, and (iii) the presence or nature of surface ligands. CeO₂ particles have a ζ potential of −22.5 ± 0.9 mV and a BET surface of 81.606 m² g⁻¹. They were functionalized with nitrilotriacetic acid (NTA) to achieve better dispersibility in aqueous solution. This is an important prerequisite for achieving homogeneous surface coating.

The rate of reaction (ROR) was determined to relate to the conversion rate to the available surface area (eqn (2)).⁴⁸ The maximum rate v_max = 0.021 μM min⁻¹ and the specific surface gave a reaction of ROR(CeO₂) = 0.0087 μmol m⁻² min⁻¹.

CeO₂ nanoparticles catalyze the oxidative bromination of N-(3-oxo-acyl)homoserine lactones (AHLs) at the activated C2 atom, forming mono- and dibromo-3-oxo-dodecanoylhomoserine lactones. The bromination is complete after approx. 60 min. The brominated AHLs are hydrolyzed in the subsequent step, the hydrolysis being complete after approx. 5 h, as demonstrated by inductively coupled plasma mass spectrometry (ICP-MS). Pseudomonas Aeroginosa bacteria use 3-oxo-dodecanoylhomoserine lactone to communicate with each other and to determine their cell count (quorum sensing, QS),²⁰ where biofilm formation is under control of QS through the production of the virulence factor pyocyanin in response to the haloperoxidase-like activity (Fig. 6, vide infra and ESI†).

Coating polycarbonate with ceria nanoparticles

Polycarbonate has a hydrophobic surface, which leads to a small surface energy. The surface was treated in an oxygen plasma. Fig. 3 shows the contact angle of untreated and oxygen plasma-treated polycarbonate. The contact angle of the pristine polycarbonate surface was 86.75° and decreased to 26.88° after a 20 min oxygen plasma treatment, i.e., the surface became significantly more hydrophilic. Hofrichter et al.⁴⁹ showed by XPS spectroscopy that plasma treatment increases the oxygen content and decreases the carbon content by generating oxygen-containing groups such as –(C–O)–, –(C=O)–, and –(O–C=O)– on the polycarbonate surface. The binding of ceria nanoparticles to polycarbonate occurs by the ester condensation of their surface hydroxyl groups with carboxylate surface groups or by hydrogen bonding between the polar groups on the polymer surface and the ceria particles.^50,51

Fig. 3 Contact angle of a water drop on (A) untreated (B) O₂ plasma-treated, and (C) CeO₂-coated polycarbonate with an optical microscopy image of the polycarbonate plates (1 × 1 cm).

The smaller contact angle of 26.88° shows a significantly higher hydrophilic surface, which facilitates the contact of the suspension with the surface of the polycarbonate and thus enables the coating with ceria nanoparticles. For the dip coating process, oxygen plasma-treated polycarbonate slides were treated in aqueous ceria suspensions and slowly pulled out of the dispersion to obtain a homogeneous coating. After dip coating, the plates were annealed at 110 °C for 16 h. Excess CeO₂ was carefully washed off before further testing. Due to the hydrophilicity of ceria on the surface, the contact angle after the coating process showed a 12.74° lower value than that of the untreated polycarbonate sample, i.e., the coating with CeO₂ nanorods increased the hydrophilicity of the polycarbonate surface. The ceria content at the surface was determined by ICPMS (1.6 μg cm⁻²). Leaching tests showed no trace of ceria in the water supernatant after 7 days. Fig. 3 demonstrates an image of untreated, plasma-treated, and CeO₂-coated PC, where there is no visual difference. In addition, this assumption is proven by spectroscopic investigation using UV/Vis spectroscopy (Fig. S9†). The polycarbonate absorbs light in the UV range for all the three samples.

The surface morphology of the untreated and CeO₂-coated polycarbonate was characterized after 10 days of storage at 7 °C by atomic force microscopy (Park System NX10, Fig. 4). When comparing the height images, the untreated polycarbonate showed a smooth topography with a low surface roughness of R_q = 1.5 nm; the polycarbonate surface coated with cerium oxide nanorods exhibited a slightly higher roughness of R_q = 5.8 nm.

Fig. 4 Surface morphology analyzed by AFM. Height images of (A) untreated polycarbonate slides, (B) polycarbonate slides coated with CeO₂ nanorods, and (C+D) phase images of CeO₂-coated polycarbonate slides.

The size of the particles can be inferred from the individual profiles.⁵² With a typical length of 80–200 nm (Fig. S4, ESI†) and an apparent typical thickness of 27–47 nm (see Fig. S3† red arrows), the CeO₂ nanorods appear to have a lower aspect ratio compared to the TEM images (Fig. 1). However, this difference originating from the tip convolution effects is bypassed when comparing the measured z-heights. Typical thicknesses of 6.2–12.7 nm (see green arrows, Fig. S3†) well fit with the observed aspect ratio. The CeO₂ nanoparticles show a distinct contrast to the polycarbonate substrate in the phase (Fig. 4C and D). The phase image of the CeO₂-coated polycarbonate plates shows the homogeneous distribution of the nanorods (Fig. 4B–D) compared to the uncoated polycarbonate slides.

The surface was also examined by scanning electron microscopy (SEM) (Fig. S1†). EDX mapping was conducted to analyze the elemental distribution and verify the presence of CeO₂. The surface was sputtered with a 1 nm gold layer to achieve a better contrast. As reported in the literature, Ce has a characteristic peak in the X-ray emission spectrum of L_α: 4.839 keV, which is reflected by the signal at 4.9 eV in the recorded EDX spectrum.⁵³ The low signal can be explained by the low concentration of CeO₂ on the surface. Also visible is the characteristic M_α signal of gold at 2.12 keV, which is attributed to the sputtered gold layer on the surface. Moreover, the signals of oxygen (O) at 0.53 keV (K_α) and carbon (C) at 0.28 keV (K_α) can be recognized.

CeO₂-Coated polycarbonate inhibits biofilm formation

The coated plates were tested regarding their potential to inhibit the biofilm formation of P. aeruginosa, a prominent Gram-negative bacterium, which is responsible for up to 10% of all healthcare-associated infections (HAI) in Germany as well as spoiling drinking water through its occurrence in drinking water pipes.⁵⁴ Due to its strong biofilm-forming behavior, simple surface cleaning with disinfectants is often not sufficient and gives rise to strains resistant to antibiotics.

To understand the effect of CeO₂ nanoparticles on biofilm formation of P. aeruginosa, crystal violet staining assay was performed to determine the amount of the adherent cells (Fig. 5A). After culturing the bacteria on polycarbonate for 72 h, the attached cells were stained with crystal violet solution. Biofilm formation was reduced by ∼75% on the coated PC surfaces. The thickness of the biofilms on the PC surfaces was quantified by confocal laser scanning microscopy (Fig. 5B). For this purpose, the bacterial biofilms were cultivated for 72 h at 30 °C on different PC samples placed in 24-well plates. The attached cells were then stained with SYTO9 and propidium iodide, two specific dyes staining the total cell count in the biofilm. The CeO₂-coated plates showed a significant reduction in the biofilm thickness to ∼20 μm compared to the uncoated PC surfaces, where the biofilm thickness was more than 50 μm. In addition, the biofilm on the coated plates grows in microcolonies and is not evenly distributed over the entire surface as it is on the uncoated plates.

Fig. 5 (A) Bacterial biofilm formation on non-coated and CeO₂-coated polycarbonate plates. Crystal violet staining assay of P. aeruginosa grown in lysogeny broth. The stained plates are shown in the lower part of the plots, (B) confocal laser scanning microscopy of P. aeruginosa biofilm on coated and non-coated surfaces. The non-coated plate served as the positive control. The planktonic cells were removed, and the remaining bacteria attached to the surfaces were stained with SYTO 9 and propidium iodide for 30 min at 30 °C. The effect of the CeO₂-coated plates on the cell attachment was then visualized. (C) Effect of CeO₂-coated polycarbonate surfaces versus non-coated plates on pyocyanin production in P. aeruginosa. Error bars represent the standard deviation of three independent experiments.

We analyzed whether the biofilm inhibiting effect of CeO₂ is mediated by blocking bacterial quorum sensing, a process designated as quorum quenching. In P. aeruginosa, the biosynthesis of pyocyanin, another virulence factor besides biofilm formation, is under control of quorum sensing. Therefore, we analyzed the pyocyanin production on CeO₂-coated and non-coated PC surfaces (Fig. 5C). For that purpose, the bacteria were grown for 72 h at 30 °C on uncoated and coated polycarbonate plates. The plates were then removed, and the remaining culture fluid was analyzed for pyocyanin. The CeO₂-coated polycarbonate plates reduced the pyocyanin formation of the biofilm grown on the surface by 54% compared to the uncoated plate. This indicates that quorum sensing-dependent pyocyanin formation is reduced by CeO₂ nanoparticles and lends support to the idea that CeO₂ also inhibits bacterial biofilm formation by quorum quenching. The remaining pyocyanin synthesis might be due to pelagic bacteria in the well, which are not directly exposed to the CeO₂ nanoparticles.

Assay for bacterial viability

To understand the process of biofilm colonization, it is important to analyze whether biofilm formation is reduced by killing the bacteria or by suppressing the group behavior.

Live/dead fluorescence assays were performed using SYTO 9 and propidium iodide (PI) dye. SYTO 9 stains only live cells, while PI penetrates only dead or damaged cells.⁵⁵ In this way, the number of living and dead cells present in the biofilm can be analyzed. Fig. 6 shows that the live/dead ratio does not change significantly for the CeO₂/polycarbonate plates and the uncoated reference, while the total amount of bacteria decreases.⁵⁵ In addition, the biofim-inhibiting activity of the CeO₂ nanorods was investigated using atomic force microscopy (AFM) and scanning electron microscopy (SEM) (Fig. S10†) for the P. aeruginosa cell culture.

Fig. 6 Quantification of P. aeruginosa biofilm formation on non-coated and CeO₂-coated polycarbonate plates. Blank plates served as yjr positive control. The planktonic cells were removed, and the remaining bacteria attached to the plates were stained with SYTO 9 (A) and propidium iodide (PI) (B). (C) Finally, the effect of the incorporated CeO₂ nanorods on bacterial biofilm formation was quantified by measuring the absorption at 485 nm for SYTO9 and 535 nm for propidium iodide, respectively. Error bars represent the standard deviation of three independent experiments.

Bacteria were cultured on CeO₂-coated and uncoated polycarbonate substrates in the presence of halide and H₂O₂. Fig. 7A+B show a homogeneous coating of bacteria with a size of 1–1.5 μm (Fig. S6†) on an uncoated polycarbonate substrate. Due to non-interrupted communication (QS) between the bacteria, they were able to generate an extracellular polymeric substance (EPS) matrix, subsequently forming a confluent biofilm and proliferate. In contrast, Fig. 7C shows a single isolated bacterium on the surface of the CeO₂-coated substrate. Here, the bacteria were brought to the surface with the LB medium but the interrupted QS prevented proliferation and inhibited biofilm formation because the AHLs used for interbacterial communication were modified and therefore inactivated by the CeO₂ nanozyme in the presence of the halide Br⁻ and H₂O₂ by oxidative bromination.²²

Fig. 7 Atomic force microscope (AFM) images of non-coated (A+B) and CeO₂-coated polycarbonate (C) with the P. aeruginosa culture.

Cytotoxicity of CeO₂ nanorods

A cell viability assay was made to determine the ability of the cells to maintain a state of survival in the presence of the CeO₂ nanorods. The cytotoxicity of the CeO₂ nanorods was examined in HeLa and RAW 264.7 cells using a luminescence cell viability assay. The CeO₂ particles showed no cytotoxicity after 24 h of incubation, even for concentrations up to 300 μg mL⁻¹.

The viability of yjr HeLa cells is constant at 100% after 2 and 24 h, even at high concentrations (Fig. 8). Similarly, the viability of the RAW 264.7 cells remains constant at 100% with increasing particle concentration. Thus, we have shown that CeO₂ coating is a non-cytotoxic approach to prevent the adhesion of bacteria to a polymer surface.

Fig. 8 Effect of CeO₂–NTA on the cell viability of HeLa and RAW 264.7 cells after 2 h and 24 h at different nanoparticle concentrations.

Conclusion

We have fabricated highly transparent CeO₂-coated polycarbonate plates with high wettability and high catalytic haloperoxidase activity to prevent bacterial adhesion. Multigram quantities of highly dispersible CeO₂ nanorods with diameters of 10–15 nm and lengths of 100–200 nm were synthesized from aqueous dispersions of cerium(III) hydroxide by oxidation with hydrogen peroxide in the presence of nitrilotriacetic acid (NTA) as the capping agent.

Transparent CeO₂-coated polycarbonate panels were prepared by oxygen plasma treatment of the polycarbonate plates, followed by dip coating with CeO₂/NTA dispersions. Low concentrations (as low as 1.6 μg cm⁻², corresponding to a monolayer coating of CeO₂ nanorods, analyzed by atomic force microscopy) were able to reduce P. aeruginosa biofilm formation on polycarbonate by at least 75%. Using cytotoxicity assays and live/dead assays, we demonstrated that the cerium oxide coating prevents bacterial adhesion, following a non-toxic mode of action. Only very low (milli/micromolar) concentrations of the substrates Br⁻ and H₂O₂ are required for haloperoxidase-like reactions to occur. These concentrations are comparable to bromide concentrations found in animal and plant body fluids and to H₂O₂ concentrations produced in daylight.⁴⁵ This makes CeO₂ nanorods a potent and “green” catalyst for antibacterial applications³³ since no chemicals are released into the environment other than those that occur anyway in natural metabolic processes of plants. Evidence for quorum quenching and therefore a non-biocidal mode of action was provided by analysis of the QS-dependent virulence factor pyocyanin. Microbial biofilms are ubiquitous on domestic, industrial, and other surfaces. Conventional biocides (strong oxidants, antibiotics, etc.) are increasingly problematic due to their lack of efficacy, non-targeted environmental effects, or development of resistance by microorganisms.

Experimental section

Materials and synthesis

Cerium(III) nitrate hexahydrate, nitrilotriacetic acid (99%), and sodium nitrilotriacetate (99+%) were purchased from Acros Organics. Hydrogen peroxide (35%) was purchased from Carl Roth. Ammonium hydroxide solution (25%) was obtained from VWR. All chemicals were used without further purification. Ultrapure water (18.2 MΩ cm⁻¹) was used for all experiments. Polycarbonate (1 mm thickness, 15 × 15 cm plates) was obtained from GoodFellow.

Synthesis of cerium dioxide nanorods. The synthesis was adapted from ref. 56 and modified for upscaling to multigram amounts of monodisperse CeO₂ nanorods. In two centrifuge tubes, a total 10 g of Ce(NO₃)₃·6H₂O were dissolved in 80 mL of H₂O. Subsequently, 8 mL of NH₄OH solution was added. After gentle shaking, the Ce(OH)₃ precipitate was centrifuged (3000 rmp, 10 min) to remove excess water. The colorless powder was redispersed in 500 mL of water, and preheated to 60 °C under magnetic and Ultra-Turrax® stirring (15 000 rpm with S 25 EC-T-C-18G ST homogenizer). After the suspension had turned homogenous, 1.4 g NTA and 0.7 g Na₃NTA were added. After further 15 min of continuous stirring, the grey-white suspension turned red after slowly adding 3 mL of H₂O₂ (35%). The temperature was slowly increased to 72 °C under ultra-Turrax®-stirring for 45 min. The reaction was allowed to cool to room temperature under magnetic stirring. To remove NH₄NO₃, the suspension was washed three times with H₂O (9000 rmp, 20 min).

Phenol red assay. The intrinsic haloperoxidase activity was determined using the phenol red assay. In this assay, tetrabromophenol blue (Br₄PR) was formed via the reaction of phenol red (PR) in the presence of a bromide source (KBr) and H₂O₂ in the presence of an active catalyst (CeO₂ nanorods). The reaction was monitored using an Agilent Cary 3500 spectrophotometer in the wavelength range from 350 to 700 nm at 25 °C. The baseline was measured at a constant temperature in a stirred (800 rpm) mixture of CeO₂ nanorods (36.3 μmol), KBr (50 μmol), and PR (0.1 μmol) in 2 mL ultrapure water (18.2 MΩ cm⁻¹) before the addition of H₂O₂ (6 μmol).

Lactone bromination. The degradation of the quorum sensing molecule 3-oxo-C₁₂-HSL by oxidative bromination was followed by time-dependent reverse high performance liquid chromatography coupled with electrospray ionisation-mass spectrometry. Several reaction mixtures (1 mL total volume) containing 900 μM 3-Oxo-HL, 20 mM KBr, 20 mM H₂O₂, and CeO₂ nanoparticles (200 μg mL⁻¹) were analyzed over a period of 5 h. The reactions were terminated by removing the catalyst by centrifugation and the supernatant was analyzed by mass spectrometry.

Michaelis–Menten kinetics. For the Michaelis–Menten kinetics, the amount of H₂O₂ was varied. The kinetics was performed at λ = 592 nm, the absorbance maximum of Br₄PR. For each measurement, the same stock solution of PR (50 μM) mixed with KBr (25 mM) was used. After the addition of 62.5 μL of nanoparticle suspension (1 mg mL⁻¹), the reaction was stirred (800 rpm) for 2 min for temperature equilibration (25 °C). Prior to H₂O₂ addition, the absorbance was set to zero. The kinetics were measured for 10 min. All data points were linearly fitted in the period of 2–8 min. The concentration was calculated using the Lambert–Beer law. The extinction coefficient was determined by measuring the absorbance of five different Br₄PR solutions (2 μM, 6 μM, 10 μM, 14 μM, 18 μM) in a quartz cuvette (d = 1 cm). Using the Lambert–Beer law, an extinction coefficient of 32 740 L mol⁻¹ cm⁻¹ was determined for BR₄PR (free acid) in ultra-pure water with an Agilent Cary 3500 UV-VIS spectrophotometer.

CeO₂ coating. Polycarbonate plates (1 mm thickness) with a size of 1 × 1 cm were washed with MQ water and ethanol to remove the surface contamination. To activate the surface, the plates were treated in oxygen plasma for 20 min (290 W, 0.1 mbar). The plasma-treated polycarbonate plates were dip-coated in a CeO₂ dispersion (9.5 ± 0.1 mg mL⁻¹) in MQ water with a dipping speed of 60 mm min⁻¹, held for 60 s in the dispersion, and pulled out with the same speed. The CeO₂-coated polycarbonate plate was dried in air at 110 °C for 16 h. Finally, the plates were washed with MQ water to remove excess CeO₂ from the surface.

Bacterial strains and growth conditions. P. aeruginosa PA14 was obtained from the lab of Dr Max Schobert (Technische Universität Braunschweig, Germany). P. aeruginosa was aerobically grown in LB medium [1% (w/v) NaCl; 1% (w/v) tryptone; 0.5% (w/v) yeast extract] at 30 °C. For the preparation of agar plates, 1.5% (w/v) agar was added to the medium.

Biofilm assays. For the quantification of bacterial biofilm production, a modified method of previously published protocols was used.⁵⁷ Briefly, P. aeruginosa was aerobically grown in LB medium overnight at 30 °C. The cultures were then diluted in LB in a volume of 1 mL per well of a 24-well polystyrene microtiter plate (Sarstedt, Nürnbrecht) at a final Optical Density at 600 nm (OD₆₀₀) of 0.5. In addition, KBr (Roth, Karlsruhe, Germany) and H₂O₂ (Roth, Karlsruhe, Germany) at a final concentration of 32 mM and 0.8 mM, respectively, were added to the wells. Then, the blank plates and CeO₂/polycarbonate composites were subjected. Subsequently, the microtiter plate was incubated for 72 h under gentle shaking (150 rpm) at 30 °C. H₂O₂ was added stepwise at a final concentration of 0.8 mM every 24 h. Then, the plates were rinsed with water to remove the planktonic cells. After drying for 5 min, 1 mL of 1% (w/v) crystal violet (Merck, Darmstadt) was added to the wells containing the plates. After 30 min incubation at room temperature, unbound crystal violet was removed by gently submerging the plates two times in water. The plate was then air-dried overnight at room temperature. For quantification, 1 mL of 30% (v/v) acetic acid (Roth, Karlsruhe) was added to the plates to solubilize the crystal violet from the biofilm. After 15 min of incubation at room temperature, the absorbance was quantified in a plate reader (Tecan, Salzburg) at 575 nm.

Fluorescence quantification of bacterial biofilm. P. aeruginosa was cultivated in the LB medium overnight at 30 °C. The cultures were then diluted in LB, in a volume of 1 mL per well of a 24-well polystyrene microtiter plate (Sarstedt, Nürnbrecht) at a final OD₆₀₀ of 0.5. In addition, KBr (Roth, Karlsruhe, Germany) and H₂O₂ (Roth, Karlsruhe, Germany) at a final concentration of 32 mM and 0.8 mM, respectively, were added to the wells. Next, the non-coated plates and CeO₂/polycarbonate slides were added to the wells. The microtiter plate was incubated for 72 h under gentle shaking (150 rpm) at 30 °C. H₂O₂ was added stepwise at a final concentration of 0.8 mM every 24 h. Then, the plates were rinsed with water to remove the planktonic cells. Subsequently, the plates were placed in 1 mL of a combined SYTO 9 and propidium iodide solution (Thermo Fisher, Pittsburgh, USA) and incubated for 30 min at 30 °C. Then, the plates were rinsed again with water and placed inside the well of a 24-well polystyrene microtiter plate. Finally, biofilm fluorescence was then quantified in a plate reader (Tecan, Salzburg) using a wavelength of 485 nm and 535 nm for the fluorophores SYTO 9 and PI, respectively.

Confocal laser scanning microscopy. P. aeruginosa was cultivated in the LB medium overnight at 30 °C. The cultures were then diluted in LB in a volume of 1 mL per well of a 24-well polystyrene microtiter plate (Sarstedt, Nürnbrecht) at a final OD₆₀₀ of 0.5. In addition, KBr (Roth, Karlsruhe, Germany) and H₂O₂ (Roth, Karlsruhe, Germany) at a final concentration of 32 mM and 0.8 mM, respectively, were added to the wells. The non-coated and CeO₂-coated plates were then added to the wells. The microtiter plate was then incubated for 72 h under gentle shaking (150 rpm) at 30 °C. Every 24 h, H₂O₂ was added at a final concentration of 0.8 mM. The plates were then rinsed with water to remove the planktonic cells. Afterward, the plates were placed in 1 mL of a combined SYTO 9 and propidium iodide 1 : 1000 dilution (Thermo Fisher, Pittsburgh, USA) and incubated for 30 min at 30 °C. The plates were then again rinsed with water and put on microscopy cover slides. The samples were examined on an Axio Imager 2 (Carl Zeiss, Jena, Germany). The fluorophore SYTO 9 was visualized with an excitation of 470 nm, while propidium iodide was visualized with an excitation of 558 nm. The images and the biofilm thickness were then processed and quantified using the ZEN 3.3 blue software.

Pyocyanin analyses. For pyocyanin production, an alternate method of already published protocols was used.⁵⁸ For this purpose, P. aeruginosa was cultivated in the LB medium overnight at 30 °C. The cultures were then diluted in LB in a volume of 1 mL per well of a 24-well polystyrene microtiter plate (Sarstedt, Nürnbrecht) at a final OD₆₀₀ of 0.5. In addition, KBr (Roth, Karlsruhe, Germany) and H₂O₂ (Roth, Karlsruhe, Germany) at a final concentration of 32 mM and 0.8 mM, respectively, were added to the wells. The non-coated plates and CeO₂/polycarbonate slides were then added to the wells. The microtiter plate was then incubated for 72 h under shaking conditions (350 rpm) at 37 °C. Every 24 h, H₂O₂ was added stepwise at a final concentration of 0.8 mM. Afterward, the composites were removed from the wells and the remaining supernatant was collected in a microreaction tube (Eppendorf, Germany). The clls were separated from the culture fluids via centrifugation at 16 × 1000g for 15 min. Then, the supernatant was passed through 0.22 μm filters (Merck, Darmstadt). Finally, the cell-free culture fluids were analyzed for absorption at 695 nm and therefore quantified in a plate reader (Tecan, Salzburg) for pyocyanin.

Cell culture. RAW 264.7 cells and human cervix carcinoma cells (HeLa) were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% FBS, 100 U mL⁻¹ penicillin, 100 mg mL⁻¹ streptomycin, and 2 mM glutamine. Cells were grown in a humidified incubator at 37 °C and 5% CO_2. Cell passaging and harvesting were performed using 0.25% Trypsin-EDTA for 5 min at 37 °C, 5% CO₂ (all reagents were from Thermo Fisher, Germany). Cell viability and cell count were determined using trypan blue and measuring by an automated cell counter (TC10, Bio-Rad, Germany).

Cell viability. Cell viability was determined using CellTiter-Glo® Luminescent Assay according to the manufacturer's manual (Promega, USA). After cell harvesting, HeLa or RAW264.7 cells were seeded in 96-well plates (item no.: 655083, Greiner BioOne, Austria) with a cell number of 5000 cells per well. Cells were incubated overnight at 37 °C and 5% CO₂ to enable cell detachment and afterward the medium was removed. Nanoparticle solutions were prepared in FBS-supplemented DMEM and added to the cells. After incubation for either 2 h or 24 h, the same volume of the CellTiter-Glo® Reagent was added and the luminescent signal was measured with an Inifinite M1000 plate reader (Tecan, Switzerland). Cell viability was displayed as the percentage of cell viability compared to the untreated cells.

Characterization

Electron microscopy. The samples for transmission electron microscopy (TEM) were prepared by placing a drop of dilute NP dispersion in ethanol on a carbon-coated copper grid. The TEM images were obtained with a FEI Tecnai 12 TWIN LaB6 at 120 kV together with a Gatan US1000 CCD-camera (16-bit, 2048 × 2048 pixels) using the Gatan Digital Micrograph software.

Samples for scanning electron microscopy (SEM) were prepared with carbon film pads on aluminum holders. A silicon wafer was used for the sample preparation and fixed on the carbon pad. A polycarbonate plate was fixed on the carbon pad and sputtered with gold (thickness 1 nm). The SEM images were obtained using an FEI Nova NanoSEM 630 equipped with an EDAX-Pegasus X4 M unit for EDX measurements. The acceleration voltage was set to 5 kV. The elemental composition was calculated using the EDAX Genesis Software Suite.

Powder X-ray diffraction. X-ray diffraction patterns were recorded on a STOE Stadi P diffractometer equipped with a Dectris My-then 1k detector in transmission mode using Mo Kα1 radiation. Crystalline phases were identified according to the PDF-2 database using the Bruker AXS EVA 10.0 software.

Zeta potentials. Zetapotentials and hydrodynamic radii were measured on a Malvern Zetasizer Nano using disposable capillary cells (DTS1070) and single-use polystyrene cuvettes. Data analysis was performed with a Malvern Zetasizer Software 8.01.4906.

BET surface area. BET measurements were conducted with a 3P Micro 300 gas adsorption instrument using nitrogen as the analysis gas at 77.4 K. The software 3P Surface Area & Pore Size Analyzer System 10.03.02 was used to analyze the recorded data.

Inductively coupled plasma mass spectrometry (ICPMS). To determine the ceria content of the coated polycarbonate plates, the plates were dissolved in CHCl₃, then dried at 45 °C, and incinerated at 600 °C. The ash was dissolved with sub-boiled HNO₃ (65%), diluted with ultra-pure water, and measured with iCAP™ RQ ICP-MS (Thermo Scientific™).

Atomic force microscopy. AFM measurements were conducted with an Park Systems NX10, using True Non-Contact™ mode and tapping mode. OMCL-AC160TS cantilever (k = 26 N m⁻¹, f ≈ 300 kHz) were used to image polycarbonate substrates and CeO₂ coating. For bacterial imaging, SD-R30-FM cantilevers (k = 2.8 N m⁻¹, f ≈ 75 kHz) were applied. The Park Systems XEI Software was used for analysis. All measurements were conducted at ambient conditions.

UV/Vis. UV/Vis measurements were conducted with an Agilent Cary 3500.

Plasma treatment. The plates were treated with oxygen plasma in a Plasmaprozessor 200G from TePla Technics Plasma GmbH at 0.1 mbar and 290 W for 20 min.

Contact angle measurements. The contact angle was measured with the contact angle measurement system OCA35 and the water drop was deposited by a Hamilton syringe (100 μL) with a hydrophobic needle.

Author contributions

O. J. carried out surface coatings and writing of original draft, F. P. carried out synthesis and writing of original draft, A. G. performed biofilm assays and contributed to original draft, J. O. and V. M. performed cytotoxicity tests, M. L. carried out electron microscopy, M. v. d. A and B. M. carried out ICP/MS analysis, A. K. performed AFM analysis, A. K., R. H. and WT conceived and designed research and revised the manuscript. All authors have given approval to the final version of the manuscript.

Conflicts of interest

The authors declare no competing financial interests.

Acknowledgements

This research was funded by a grant from the Deutsche Forschungsgemeinschaft (DFG) to W. T. (grant number TR 210/38-1) and the Bundesministerium für Bildung und Forschung (BMBF) Neue antimikrobielle Wirkstoffe gegen Biofilme (project number 031B0583A) to R. H. and Entwicklung innovativer biozider Nanopartikel zur Anwendung in der Kunststofftechnik (project number 19585N) to W. T. We thank S. Berinskat and F. Ludwig for X-ray diffraction measurements and Prof. A. Möller for access to the diffractometer. We are indebted to Prof. Rudolf Zentel (Department Chemie (JGU Mainz)) for the access to the Zetasizer. Thanks also apply to Prof. Hans-Jürgen Butt (MPI for Polymer Research Mainz) for providing the contact angle measurement system and the plasma cleaner. The microscopy equipment is operated by the Electron Microscopy Center (EMZM). Confocal laser scanning microscopy was performed at the Institute for Biotechnology and Drug Research (IBWF) Mainz.

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Footnotes

† Electronic supplementary information (ESI) available: Fig. S1 demonstrates the scanning electron microscopy (SEM) image (A) and energy-despersive X-ray (EDX) spectrum (B) of the CeO₂-coated surface of polycarbonate. This is followed by information about the uncoated and CeO₂-coated polycarbonate surface investigated by AFM (Fig. S2–4). The uncoated and CeO₂-coated PC surface with P. aeruginosa were also examined by AFM (Fig. S5+6). Fig. S7 demonstrates the microscope images of HeLa and RAW cells on the surface of CeO₂-coated polycarbonate. In conclusion, an image of aqueous CeO₂-suspersion is shown in Fig. S8. Fig. S9 shows the UV/Vis spectra of uncoated, plasma-treated, and CeO₂-coated PC, showing no visual difference after coating. See DOI: 10.1039/d1nr03320d

‡ These authors contributed equally to this work.

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