# Boosting Membrane Interactions and Antimicrobial Effects of Photocatalytic Titanium Dioxide Nanoparticles by Peptide Coating

Lucrezia Caselli, Elisa Parra-Ortiz, Samantha Micciulla, Maximilian W.A.

Skoda, Sara Malekkhaiat Häffner, Emilie Marie Nielsen, Mariena J.A. van der Plas, Martin Malmsten

► To cite this version:

Lucrezia Caselli, Elisa Parra-Ortiz, Samantha Micciulla, Maximilian W.A. Skoda, Sara Malekkhaiat Häffner, et al.. Boosting Membrane Interactions and Antimicrobial Effects of Photocatalytic Titanium Dioxide Nanoparticles by Peptide Coating. Small, 2024, 20 (30), pp.2309496. 10.1002/smll.202309496 . hal-05330160

HAL Id: hal-05330160

https://cnrs.hal.science/hal-05330160v1

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# Boosting Membrane Interactions and Antimicrobial Effects of Photocatalytic Titanium Dioxide Nanoparticles by Peptide Coating

Lucrezia Caselli, Elisa Parra-Ortiz, Samantha Micciulla, Maximilian W. A. Skoda, Sara Malekkhaiat Häffner, Emilie Marie Nielsen, Mariena J.A. van der Plas, and Martin Malmsten*

Photocatalytic nanoparticles offer antimicrobial effects under illumination due to the formation of reactive oxygen species (ROS), capable of degrading bacterial membranes. ROS may, however, also degrade human cell membranes and trigger toxicity. Since antimicrobial peptides (AMPs) may display excellent selectivity between human cells and bacteria, these may offer opportunities to effectively “target” nanoparticles to bacterial membranes for increased selectivity. Investigating this, photocatalytic TiO₂ nanoparticles (NPs) are coated with the AMP LL-37, and ROS generation is found by C₁₁-BODIPY to be essentially unaffected after AMP coating. Furthermore, peptide-coated TiO₂ NPs retain their positive ζ-potential also after 1–2 h of UV illumination, showing peptide degradation to be sufficiently limited to allow peptide-mediated targeting. In line with this, quartz crystal microbalance measurements show peptide coating to promote membrane binding of TiO₂ NPs, particularly so for bacteria-like anionic and cholesterol-void membranes. As a result, membrane degradation during illumination is strongly promoted for such membranes, but not so for mammalian-like membranes. The mechanisms of these effects are elucidated by neutron reflectometry. Analogously, LL-37 coating promoted membrane rupture by TiO₂ NPs for Gram-negative and Gram-positive bacteria, but not for human monocytes. These findings demonstrate that AMP coating may selectively boost the antimicrobial effects of photocatalytic NPs.

## 1. Introduction

Due to growing antibiotic resistance, there is a need for new types of antimicrobial agents.[1,2] Nanoparticles (NPs) are currently receiving considerable interest in this context, as many such materials display antimicrobial effects involving, e.g., direct membrane destabilization, oxidative degradation of essential bacteria components, and heating-induced lysis.[3–5] Among these, photocatalytic nanomaterials display potent antimicrobial effects under illumination. In photocatalysis, free electrons and electron holes are formed on light excitation. These may react with water or dissolved oxygen to form reactive oxygen species (ROS), which, in turn, trigger antimicrobial effects by oxidizing membrane lipids and other essential bacterial components.[6–8]

There are many types of photocatalytic nanomaterials, including TiO₂, and quantum dots, as well as various carbon-based photocatalysts, such as fullerenes and graphitic carbon nitride nanosheets.[9]

L. Caselli, E. Parra-Ortiz, S. M. Häffner, E. M. Nielsen, M. J. van der Plas, M. Malmsten
Department of Pharmacy
University of Copenhagen
Copenhagen DK-2100, Denmark
E-mail: martin.malmsten@sund.ku.dk

S. Micciulla
Institut Laue-Langevin
CS 20156, Grenoble Cedex 9 38042, France

S. Micciulla
Laboratoire Interdisciplinaire de Physique (LIPhy)
Saint Martin d'Hères 38402, France

M. W. A. Skoda
ISIS Pulsed Neutron and Muon Source
Rutherford Appleton Laboratory
Harwell OX11 0QX, UK

L. Caselli, M. Malmsten
Department of Physical Chemistry 1
Lund University
Lund SE-22100, Sweden

<figure><img src="image_3.png" /></figure>

The ORCID identification number(s) for the author(s) of this article can be found under https://doi.org/10.1002/smll.202309496

© 2024 The Authors. Small published by Wiley-VCH GmbH. This is an open access article under the terms of the Creative Commons Attribution License, which permits use, distribution and reproduction in any medium, provided the original work is properly cited.

DOI: 10.1002/smll.202309496
Among these, TiO₂ NPs have received particularly intense interest. Photocatalytic effects of TiO₂ NPs have been found to depend, e.g., on crystal structure and morphology.[10] In addition, photocatalytic effects of TiO₂ can be boosted through doping and heterojunction formation, aiming at i) decreasing the band gap (making the nanoparticles excitable by visible light), ii) increasing free radical formation, and iii) decreasing the rate of charge re-combination. Investigating antimicrobial effects of such systems (further examples can be found in),[9,10] Arora et al. found potent antimicrobial effects of TiO₂ NPs against multi-drug resistant (MDR) *P. aeruginosa* under UV illumination.[11] Similarly, Ahmed et al. investigated antimicrobial effects of TiO₂ NPs on 25 *P. aeruginosa* isolates that were highly or completely resistant to antibiotics alone and found antimicrobial effects of TiO₂ NPs for all MDR isolates. Furthermore, TiO₂ NPs combined with the antibiotics ciprofloxacin, cefepime, amikacin, or ceftriaxone showed synergistic activity against all MDR isolates.[12] Illustrating the role of TiO₂ structure on such effects, Molina-Reyes et al. compared ROS generation and antimicrobial effects of TiO₂ nanoparticles and nanotubes against *E. coli*. While ROS generation was comparable for these systems, the antibacterial activity was higher for TiO₂ nanotubes, an effect of the latter allowing closer surface contact with bacteria.[13]

Despite potent antimicrobial effects thus displayed by TiO₂ and other photocatalytic nanomaterials, key challenges in their development toward antimicrobial therapeutics include their toxicity, as well as a currently poor understanding of their antimicrobial spectrum width. Illustrating the latter, Sułek et al. found a 7-log reduction of Gram-positive *Staphylococcus aureus* (*S. aureus*) under UV illumination in the presence of porphyrin-doped TiO₂ NPs, but a much poorer effect against Gram-negative *Escherichia coli* (*E. coli*).[14] In contrast, Gao et al. found Fe³⁺-doped g-C₃N₄ to be equally potent against Gram-negative *Pseudomonas aeruginosa* (*P. aeruginosa*), Gram-negative *E. coli*, and Gram-positive *S. aureus*,[15] whereas Yadav et al. found antimicrobial effects of g-C₃N₄ nanosheets to be stronger for *E. coli* than for *S. aureus*.[16] Considering this scatter in functional performance, further work is clearly needed to clarify how antimicrobial effects and toxicity of photocatalytic nanomaterials depend on differences in membrane structure and composition. Composition-wise, cholesterol is a key component (up to 45% mol) in human cell plasma membranes, but it is not present in bacteria.[17,18] Furthermore, whereas human cell membranes are rich in zwitterionic phospholipids (e.g., phosphatidylcholine, PC), bacterial membranes contain large fractions of anionic phospholipids (e.g., phosphatidylglycerol, PG); ≈25% mol), despite some minor variations between bacterial strains.[18–20] Such systems are

therefore frequently used as model systems for cell and bacteria membranes, respectively.[3,9]

Due to its large band gap,[21] TiO₂ requires illumination in the UV spectrum to display photocatalytic properties. While this may be a disadvantage in practical applications due to limited tissue penetration of UV light, it makes TiO₂ NPs suitable for mechanistic studies, since “in darkness” measurements can be achieved under normal laboratory lighting, conveniently “turning on” ROS generation by UV illumination. Previously, we reported on the influence of lipid composition on membrane susceptibility to photocatalytic degradation by bare TiO₂ NPs. Whereas cholesterol was found to provide stabilization against TiO₂-mediated oxidation of phosphatidylcholine-based bilayers, anionic phosphatidylglycerol loading increased susceptibility to oxidative destabilization.[22,23] Although such compositional differences thus provide some selectivity between membranes mimicking those of bacteria and human cells, the development of photocatalytic nanomaterials toward antimicrobial therapeutics requires further strengthening of such selectivity. In this context, antimicrobial peptides (AMPs) provide interesting opportunities. AMPs play a key role in innate immunity, and reach antimicrobial potency by several action mechanisms, the most important of which being the physical destabilization of bacterial membranes.[24,25] In addition to such direct destabilization of bacterial membranes, some AMPs interact with bacterial lipopolysaccharides to suppress inflammation and subsequent uncontrolled host response to the infection.[26,27] A key aspect of AMPs is that these can be broadly potent against bacteria, including those resistant to antibiotics, yet display low toxicity against human cells.[28,29] Considering this, AMPs may potentially be used as a surface coating to effectively “target” nanomaterials to bacterial cells. In this study, we therefore investigate if TiO₂ NPs can be coated with AMPs to boost selectivity between bacteria and cell membranes further. In doing so, however, several key questions must be addressed: i) Considering that ROS generation depends on photogenerated electrons and holes reaching the TiO₂/water interface to react with water and dissolved oxygen to form ROS, how does a (potentially dense) peptide layer influence ROS formation? ii) Since also peptides are susceptible to photocatalytic oxidation, can a window be found, in which the adsorbed peptide layer does not impair ROS formation to allow oxidative degradation of bacterial membranes, yet remains sufficiently stable for allowing targeting of these by the peptide-coated nanoparticles? iii) Since membrane lysis of AMPs depends on their ability to orient and insert into lipid membranes, properties which may be suppressed when AMPs bind to the nanoparticle surface, how do these effects influence the destabilization of lipid membranes?

In addressing these questions, we employed a battery of methodologies previously established by us for investigating photocatalytic degradation of phospholipid membranes by bare TiO₂ NPs, including neutron reflectometry (NR), quartz crystal microbalance with dissipation monitoring (QCM-d), light scattering, and fluorescence spectroscopy. Based on considerations of bacteria and cell membrane composition, we selected model bilayers based on PC, supplemented with either PG (+PG) or cholesterol (+Chol) to mimic bacterial or mammalian membranes, respectively. PC lipid membranes, devoid of PG or cholesterol, were also included in the study as reference samples.
<figure><img src="image_4.png" /><figcaption>**Figure 1.** A) ζ-potential and number average particle size of TiO₂ NPs as a function of pH for 100 ppm TiO₂ in 10 mM acetate pH 3.4 and pH 5.4 or 10 mM Tris pH 7.4 and 9.4. B) Corresponding results for 100 ppm TiO₂ NPs loaded at varying concentrations of LL-37 in 10 mM acetate, pH 5.4. C) ζ-potential of TiO₂ NPs (100 ppm) coated with LL-37 (10 µM) in 10 mM acetate, pH 5.4, before and after UV illumination for 1 and 2 h. Measurements were performed in triplicate.</figcaption></figure>

Regarding the choice of specific phospholipid acyl chains, palmitoyloleoyl PC and PG were selected for relevance in bacterial and cell membranes, but also for forming largely defect-free supported bilayers, necessary for neutron reflectometry experiments. To understand how the physicochemical results thus obtained for model lipid membranes translate to antimicrobial effects and cell toxicity, in vitro effects of AMP-coated TiO₂ NPs on bacteria and cells were also studied by confocal microscopy and cell toxicity assays. Since photocatalytic nanoparticles are particularly interesting for combatting superficial infections, these studies were performed at slightly acidic and neutral conditions, relevant for the pH range of skin.[30,31]

## 2. Experimental Section

### 2.1. Materials

TiO₂ NPs (anatase, 4–8 nm) were from PlasmaChem GmHb (Berlin, Germany). Particle size distributions were checked with cryo-TEM and Small-Angle X-ray scattering (SAXS). To obtain reliable information on the primary particles (i.e., not affected by aggregation), measurements were performed in 10 mM Acetate buffer, pH 3.4, where TiO₂ NPs show optimal colloidal stability (Figure 1A). SAXS results were recorded in the 0.007–0.56 Å⁻¹ q-range and fitted to a model function from the Igor-Pro-based NCNR SANS package,[32] accounting for a polydisperse population of spheres with uniform scattering length density (SLD) and a logNormal size distribution (Figure S1A, Supporting Information). The model yielded an average nanoparticle size of 2 ±1 nm, with a polydispersity index (PDI) of

0.5. The SAXS profile was also analyzed using Guinier model-free analysis[33,34] (Figure S1B, Supporting Information), performed in the low-q range (0.007-0.04 Å⁻¹), yielding similar results (average diameter 3 ± 1 nm). In line with the SAXS analysis, cryo-TEM results (Figure S1C, Supporting Information) showed the occurrence of polydisperse particles with an average diameter of 2 ± 1 nm, but sizes spanning the entire range of 0–10 nm. LL-37 (LLGDFFRKSKEKIGKEFKRIVQRIKD-FLRNLVPRTES) was obtained from Thermo Fisher Scientific (Waltham, Massachusetts, USA), and was of > 96% purity. Poly(arginine) (Mw 5–10 kDa), Tris buffer (Trizma base, ≥99.9%), acetic acid (99.7%), sodium acetate (≥99%), and lipopolysaccharides from *Escherichia coli* O111:B4 (L2630) were from Sigma Aldrich (St. Luis, USA). Palmitoyloleoylphosphoglycerol (POPG), palmitoyloleoylphosphocholine (POPC), palmitoyl-arachidonoyl phosphocholine (PAPC), and cholesterol (bovine, Chol) (all >99%) were from Avanti Polar Lipids (Alabaster, US). C₁₁-BODIPY 581/591 was from Molecular Probes/Thermo Fisher Scientific (Waltham, Massachusetts, USA). Ultra-pure MilliQ water (MQ, 18.2 MΩ·cm) and D₂O (99% deuterated, Sigma Aldrich (St. Luis, USA)) were used throughout. All other chemicals were of analytical grade.

### 2.2. Dry Lipid Film and Liposome Preparation

Lipid films and liposomes were prepared as described previously.[22] In short, lipids were dissolved in either chloroform or chloroform/methanol (3:1 v/v mixture for dissolving POG) to form the following lipid mixture: 75/25 POPC/PAPC
("PC"), 50/25/25 POPC/PAPC/POPG ("+PG"), or 65/25/10
POPC/PAPC/Chol ("+Chol") mixtures (molar ratios). The final
lipid concentration was 10 mg mL⁻¹. Note that the inclusion
of PAPC was not critically important for oxidative degradation,
but still provides an advantage by reducing the UV exposure
dose required, and hence also the risk for sample heating. The
solvent was then thoroughly removed, first under N₂ flow and
then under vacuum overnight. The lipid films were re-hydrated
to 1 mg mL⁻¹ with MQ water for liposomes for supported lipid
bilayers (SLB), and with buffer for liposomes to be investigated
in suspension. The solution was vortexed and bath-sonicated
for 30 min to obtain multilamellar vesicles (MLVs). Small unil-
amellar vesicles (SUVs) for supported bilayers were formed
by 15 min of tip sonication (UP50H, Hielscher Ultrasonics
GmbH, Germany), which gave SUVs of ≈30 nm diameter. Large
unilamellar vesicles (LUVs) were formed by vesicles extrusion
through polycarbonate filters (100 nm pores) using a LipoFast
miniextruder (Avestin, Ottawa, Canada), to be used for ROS
generation studies.

2.3. Size and ζ-Potential Measurements

Dynamic and electrophoretic light scattering (DLS and ELS; 173° back-scattering angle) was performed using a Zetasizer Nano ZSP (Malvern Pananalytical Ltd., Malvern, UK) to obtain particle size and ζ-potential characteristics. Measurements consisted of 10 runs of 10 s each, using automatic attenuation, are reported as the number-average of the effective particle diameters, and were performed in triplicate at 25 °C. Apart from bare and LL-37-coated TiO₂ NPs, such measurements were performed also for LL-37-coated non-photocatalytic mesoporous silica nanoparticles (160±1 nm diameter)[³⁵] to investigate the effects of UV exposure as such on peptide oxidation (Figure S3, Supporting Information).

2.4. Small Angle X-Ray Scattering (SAXS)

Structural features of bare TiO₂ NPs were studied using the
laboratory-based Xeuss 3.0 instrument (Xenocs, France) at The
Center for Scattering Methods (CSM) in the Faculty of Science,
Lund University (Sweden), equipped with an X-ray source pro-
ducing a photon beam with a wavelength of 1.34 Å. The scatter-
ing patterns were recorded with an Eiger2 R 1 M 2 D-detector
(Dectris) and azimuthally integrated using the XSACT program
available with the equipment creating 1D scattering curves. The
radially averaged intensity, I(q), was given as a function of the
scattering vector q = 4πsinθ/λ, where λ is the wavelength and 2θ
was the scattering angle. For the measurements, a dispersion of
5000 ppm bare TiO₂ NPs in 10 mM Acetate buffer, pH 3.4, was
sealed in a glass capillary and the scattering profile was acquired
at 25 °C.

**2.5. Cryogenic Transmission Electron Microscopy (cryo-TEM)**

Cryo-TEM experiments were performed on a JEM-2200FS trans-
mission electron microscope (JEOL) at the National Center for

High-Resolution Electron Microscopy (nCHREM), Lund Univer-
sity. Zero-loss images were recorded at 200 kV on a bottom-
mounted TemCam-F416 camera (TVIPS) using SerialEM un-
der low-dose conditions. Specimens were prepared using an au-
tomatic plunge system (Leica EM GP) with the environmental
chamber at 20 °C and 90% relative humidity. Four mL droplets
were deposited on a carbon-coated grid (Ted Pella) and blotted
with filter paper to remove excess fluid. The grid was plunged into
liquid ethane (–184 °C) to ensure rapid vitrification. Specimens
were then stored in liquid nitrogen (–196 °C) until transferred
into the microscope using a cryo transfer tomography holder
(Fischione Model 2550). Cryo-TEM images for 5000 ppm bare
TiO2 NPs in Acetate buffer, pH 3.4, were acquired at different
magnifications, and particle size histograms were obtained from
cryo-TEM images using ImageJ (National Institutes of Health,
Bethesda, USA).[36,37] Weibull fits[38] of particle size histograms
were performed to obtain average particle size and standard de-
viation.

2.6. C₁₁-BODIPY Oxidation Assay

C₁₁-BODIPY experiments were performed as described in de-
tail elsewhere.[22] In brief, 0.5 mol% of the probe was added be-
fore lipid film drying under Ar atmosphere. After hydration and
extrusion, 0.5 mg mL⁻¹ LUVs were illuminated by UV in situ
(254 nm; 3 mW cm⁻²) in the presence or absence of either bare
or peptide-coated TiO₂ NPs (100 ppm). Fluorescence spectra were
acquired, and oxidation was quantified by monitoring the spec-
tral shift from red (λmax = 594 nm) to green (λmax = 520 nm).
These experiments were also performed in the presence of ROS
scavengers, i.e., D-mannitol (*OH scavenger*)[39] and SOD (*O₂*⁻
scavenger),[40] to obtain information on the role of different ROS
on lipid oxidation. All measurements were performed in dupli-
cate at 37 °C.

2.7. Quartz Crystal Microbalance with Dissipation Monitoring
(QCM-d)

QCM-d measurements were performed as described in detail
before.[22] In short, these measurements were performed us-
ing a QSense Analyzer equipped with both standard and UV-
transparent window modules (Biolin Scientific, Sweden). Sup-
ported bilayers were prepared by SUVs' deposition on silicon
dioxide surfaces (QSense QSX 303 SiO₂, 4.95 ± 0.05 MHz). For
this, sample cells and tubing were cleaned with 2% Hellmanex
and multiple MQ rinses under bath sonication, followed by rins-
ing with ethanol and N₂ drying. After similar cleaning, the SiO₂
surfaces were plasma cleaned (PDC-32G, Harrick Plasma, USA)
for 2 min before cell assembly. SUVs (0.1 mg mL⁻¹ in MQ)
were injected into the chamber and their deposition, rupture, and
full bilayer formation were monitored from the variation of fre-
quency shift and dissipation changes (ΔF and ΔD, respectively).
After rinsing with MQ water to remove non-adsorbed vesicles,
the chamber was flushed with a buffer solution containing ei-
ther bare or peptide-coated TiO₂ NPs (100 ppm). Subsequently,
samples were exposed to UV light at a distance of 6 cm (Spec-
troline ENF-260C, 254 nm; 3 mW cm⁻²) to trigger oxidation. All
measurements were performed at 37 °C at least in triplicate.
## 2.8. Neutron Reflectometry (NR)

NR experiments were performed as described in detail previously.[22] In brief, experiments on +PG bilayers interacting with bare and LL-37-coated TiO₂ NPs were performed on the D17 vertical reflectometer (Institute Laue-Langevin, Grenoble, France), operating in time-of-flight mode.[41,42] The Q-region of interest (0.01 to 0.3 Å⁻¹) was accessed using two incident angles (0.8° and 3.0°), wavelength range between 2 and 30 Å and wavelength resolution between 1 and 4%. For kinetic measurements, an intermediate angle of 1.8° was used to monitor the Q-region where the main changes in the reflectivity profile were expected to occur, and a divergent beam geometry was set to achieve improved statistics with shorter acquisition times (60 s). Experiments on PC bilayers interacting with LL-37-coated TiO₂ NPs were performed using the OffSpec reflectometer (ISIS Pulsed Neutron and Muon Source, Rutherford Appleton Laboratory, Harwell, UK).[43,44] Three incident angles (0.3°, 1.0° and 2.3°) were used to cover the Q-region from ≈0.01 to 0.35 Å⁻¹. For kinetic measurements, the reflectometry changes at the first two angles were recorded every 10 min, acquiring data for 30 s at 0.3° and for 9 min 30 s at 1.0°. Solid-liquid flow cells, the top plate of which was modified with a 30 mm diameter circular opening to allow in situ UV irradiation were used, together with UV-transparent quartz blocks (80×50×15 mm, 1 face polished, RMS < 4.5 Å, PI-KEM Ltd., Tamworth, UK). The blocks were cleaned in dilute acid piranha solution (5/4/1 v/v H₂O/H₂SO₄/H₂O₂) at 80 °C for 15 min, rinsed thoroughly in MQ, followed by 10 min of UV-ozone treatment (UV/Ozone ProCleaner, BioForce Nanosciences, USA). HPLC tubing, PEEK troughs, and O-rings were cleaned with a 2% Hellmanex solution (Hellma Analytics, UK) in bath sonication, then both thoroughly rinsed and sonicated in MQ. A circulating water bath was used during the measurements to keep the sample cells at 37 °C.

The bare quartz surfaces were first characterized in D₂O and MQ. Then, tip-sonicated SUVs (0.1 mg mL⁻¹) were injected by syringe and allowed to deposit for 20 min, after which the excess was rinsed off with 10 mL MQ and 10 mL of the desired buffer at 2 mL min⁻¹. The bilayers thus formed were characterized in two contrasts, 10 mM acetate in MQ (HAc) or in D₂O (DAC), before treatment. Contrast exchange was performed by pumping 20 mL of the desired buffer at 2 mL min⁻¹, all buffers being degassed by bath sonication prior to experiments. Samples were incubated for 10 min with bare or LL-37-coated TiO₂ NPs (100 ppm) in DAC at the desired pH. The bilayers were then subjected to in situ UV irradiation (Spectroline ENF-260C, 254 nm; 3 mW cm⁻²) for 2 h. During this time, kinetic measurements were taken every minute in the middle-Q region as mentioned above. Immediately after UV exposure, the whole Q-range was measured in DAC, after which the bilayer was rinsed and characterized in both contrasts. Experimental NR profiles were fitted by using the Genetic Optimization method available on the Motofit software within the analysis package IGOR Pro.[45,46] A series of parallel layers were used to model the interfacial structure, each of these described by a set of physical parameters, including thickness, roughness, hydration, and neutron scattering length density (SLD). The best fits of these parameters were then converted into SLD profiles, which represent the density distribution in the direction perpendicular to the reflecting interface. A Monte Carlo error analysis

allowing for refitting data 200 times was employed to minimize the uncertainty associated with data fitting.[47]

## 2.9. Live/Dead Bacterial Viability Assay

E. coli ATCC 25922 and S. aureus ATCC 29213 were stained using the LIVE/DEAD BacLight Bacterial Viability Kit (Thermo Fisher Scientific Inc., Waltham, USA). Bacteria were grown to stationary phase in 25 mL Lennox broth (LB Broth; Sigma Aldrich (St. Luis, USA)) overnight at room temperature, shaking at 180 rpm. The bacteria were pelleted and washed by centrifugation (10 000 × g, 10 min, twice) and re-suspended in 10 mM acetate buffer, pH 5.4. Nanoparticle dispersions, either bare or loaded with LL-37 at their maximum loading capacity, were added to bacteria for a final TiO₂ NPs concentration of 100 ppm (10 µM LL-37) and an optical density OD₆₀₀ of 1.2, corresponding to 12 × 10⁸ colony forming units (CFU) mL⁻¹ for E. coli and 6 × 10⁸ (CFU) mL⁻¹ for S. aureus (confirmed on subsequent dilution, plating, culture, and counting of colonies), in the final mixture. Samples were then placed in quartz cuvettes and incubated for 1 hour at room temperature, either in darkness or in the presence of UV illumination (Spectroline ENF-260C, 254 nm; 3 mW/cm²), the latter at a cuvette-lamp distance of 6 cm. Samples were subsequently diluted in 10 mM acetate buffer, pH 5.4, to obtain an OD₆₀₀ of 0.6, corresponding to 6 × 10⁸ (CFU) mL⁻¹ for E. coli and 3 × 10⁸ (CFU) mL⁻¹ for S. aureus. This was followed by 10 min of staining of 200 µL of sample with 0.5 µL of a 1/1 (v/v) mixture of the fluorescent probes SYTO 9 (excitation/emission maxima 480/500 nm) and propidium iodide (excitation/emission maxima 490/635 nm).[48] Subsequently, samples were imaged through confocal microscopy.

## 2.10. Confocal Microscopy

Bacteria were plated onto a cover slide at 10⁸ CFU mL⁻¹ and then imaged with a 100×/1.25 oil objective using a Leica DMi8 confocal microscope (Leica Microsystems, Washington, D.C., USA). For each sample, 20 randomized, wide-field images (100 µm × 100 µm) were collected. Quantification of the fraction of Live/Dead bacteria was performed with the software ImageJ (National Institutes of Health, Bethesda, USA).[36,37] Experiments were run in triplicate at 25 °C.

## 2.11. Lactate Dehydrogenase (LDH) Assay

To investigate cell toxicity, THP1 monocytes (THP1-Xblue-CD14 reporter cells (InvivoGen, France) were cultured according to the manufacturer's instructions, and nanoparticle-induced release of the intracellular enzyme LDH was monitored. For this,

Hundred ppm bare or LL-37-coated TiO₂ NPs were mixed with 100 ppm (0.1 mg mL⁻¹) lipopolysaccharide (LPS) from E. coli O111:B4 in 10 mM Tris, pH 7.4. Samples were then placed in quartz cuvettes and incubated for 2 hours at room temperature, either in darkness or in the presence of UV illumination (Spectroline ENF-260C, 254 nm; 3 mW/cm²), at a cuvette-lamp distance of 6 cm. Next, cells (1 × 10⁶ cells mL⁻¹) were incubated with LPS/bare TiO₂ or LPS/LL-37-coated TiO₂. To be able to compare
results for cells and bacteria, LDH experiments were performed at an average of 250 TiO₂ NPs per cell, which is comparable to the range used for the confocal microscopy studies of bacteria, the latter being in the range of 200 (E. coli) to 400 (S. aureus) nanoparticles per CFU. After 22–24 h incubation at 37 °C in 5% CO₂, LDH release was measured in triplicate using a lactate dehydrogenase assay kit (Invitrogen CyQUANT LDH Cytotoxicity Assay, Fisher Scientific), according to manufacturer's instructions.

## 2.12. Statistical Analysis

All data were presented as means with standard errors. For each experiment (except for neutron reflectometry experiments), measurements were performed at least in triplicate. Errors associated with the bilayer's structural parameters from neutron reflectometry were obtained through a Monte Carlo error analysis,[47] available on the Motofit software within the analysis package IGOR Pro,[45,46] which allowed for refitting experimental NR profiles 200 times, to minimize the uncertainty associated to data fitting.

# 3. Results

## 3.1. Peptide Coating of TiO₂ NPs

TiO₂ NPs have previously been reported to display pH-dependent charge density, with an isoelectric point at pH 6–6.5.[49,50] In line with this, TiO₂ NPs were found to display a net positive ζ-potential at pH 3.4 and 5.4, and a negative charge at pH 7.4 and 9.4 (Figure 1A).[23] Mirroring this, TiO₂ nanoparticles displayed pronounced aggregation close to the isoelectric point, while much less so at pH 3.4 and 9.4. By loading TiO₂ NPs with LL-37 with increasing peptide concentrations at pH 5.4, an increasingly positive ζ-potential was measured, suggesting a concentration-dependent peptide binding to the TiO₂ nanoparticle surface (Figure 1B). For TiO₂ NPs fully loaded with LL-37 (≥10 µM), the ζ-potential reaches +38 ± 2 mV, providing improved colloidal stability to the peptide-loaded nanoparticles. Furthermore, the LL-37 coating was found to be relatively resilient to UV degradation, as indicated by only partially suppressed positive ζ-potential after up to 2 h of UV illumination (Figure 1C). At pH 7.4, similar effects were observed, although a higher peptide concentration was needed to achieve saturation in positive ζ-potential and colloidal stability (Figure S2A,B, Supporting Information). In contrast to partial peptide layer degradation on prolonged UV illumination for peptide-coated TiO₂ NPs, no such effects were observed for non-photocatalytic mesoporous SiO₂ nanoparticles under these conditions (Figure S3A,B, Supporting Information), demonstrating that UV itself (i.e., in the absence of TiO₂ NPs) does not induce any significant peptide degradation or oxidation under the experimental conditions of the study.

## 3.2. ROS Generation and Lipid Oxidation

Since the adsorbed peptide layer may detrimentally influence the efficiency by which photogenerated electrons and holes reach the particle/water interface to be able to react with water and oxygen,

<figure><img src="image_5.png" /><figcaption>**Figure 2.** C₁₁-BODIPY oxidation rates for 100 ppm bare TiO₂ NPs or TiO₂ NPs coated with 10 µM LL-37 on PC (top), +Chol (middle), and +PG (bottom) LUVs exposed to in situ UV illumination for 120 min, as well as corresponding results on effects of addition of D-mannitol (*OH scavenger, 500 mM) and SOD (*O₂⁻ scavenger, +50 U mL⁻¹) in 10 mM acetate, pH 5.4. Measurements were performed in triplicate.</figcaption></figure>

key for ROS generation, we next investigated how peptide coating affected lipid oxidation in PC, +Chol and +PG large unilamellar vesicles (LUVs) by TiO₂ NPs. As shown in Figure S4 (Supporting Information), oxidation was very low (3–5% min⁻¹) in the absence of TiO₂ NPs. When bare TiO₂ NPs were present, the oxidation rate was 4–6 times higher for +PG than that for PC or +Chol, the latter showing similar oxidation rates as in the absence of nanoparticles at pH 5.4 (Figure 2; see also Figure S5 (Supporting Information) for corresponding oxidation kinetics) and 7.4 (Figures S7 and S8A, Supporting Information, for oxidation rates and oxidation kinetics, respectively). Importantly, LL-37 coating did not markedly influence ROS generation, indicating that the peptide layer was sufficiently “open” to allow photogenerated electrons and holes to reach the particle surface to form ROS. Furthermore, the addition of D-mannitol (*OH scavenger*)[39] to LUV suspensions resulted in a significant suppression of the oxidation
<figure><img src="image_6.png" /><figcaption>**Figure 3.** A) QCM-d results showing the adsorption of LL-37-TiO₂ NPs on supported PC, +Chol, and +PG bilayers in 10 mM acetate, pH 5.4. B) Adsorption of bare TiO₂ and LL-37-TiO₂ NPs on +PG bilayers in 10 mM acetate, pH 5.4. Measurements were performed in triplicate. Representative QCM-d profiles are reported in Figure S11A (Supporting Information).</figcaption></figure>

rate for bare TiO₂ NPs both at pH 5.4 (Figures 2 and S6A) and 7.4 (Figures S7 and S8B, Supporting Information), showing •OH radicals to be formed during UV illumination. In contrast, SOD (●O₂⁻ inhibitor)[40] had a limited impact on the oxidation of bare TiO₂ NPs (Figure 2 and Figure S6B, Supporting Information, for pH 5.4, and Figures S7 and S8C, Supporting Information for pH 7.4). In line with this, experiments under de-aerated conditions gave results similar to those obtained under aerated conditions (Figure S9A,B, Supporting Information), showing the effects of dissolved oxygen to be relatively minor. Analogous effects of D-mannitol and SOD were observed for LL-37-TiO₂ NPs at both pH 5.4 (Figure 2 and Figure S6A,B, Supporting Information) and pH 7.4 (Figures S7 and S8B,C, Supporting Information).

While C₁₁-BODIPY experiments provide valuable information on UV-induced ROS generation and subsequent oxidation triggered by these, these report on oxidation of the dye, incorporated into lipid bilayers, rather than on oxidation of the bilayer phospholipids. Considering this, QCM-d was also included in our investigation to directly monitor the consequences of lipid oxidation. For this, lipid bilayers from PC, +Chol, or +PG SUVs were formed on SiO₂-coated quartz crystals. The successful formation of homogeneous lipid bilayers was confirmed by changes

in frequency (ΔF ≈ −25-28 Hz) and dissipation (ΔD ≈ 0) (Figure S10, Supporting Information).[51] LL-37-coated TiO₂ NPs were found to bind to all the membranes investigated (Figure 3A and S11A for representative QCM-d profiles). Quantitatively, however, nanoparticle deposition was much higher on anionic +PG bilayers + (−66 ± 13 Hz), than on zwitterionic PC or +Chol membranes (−32 ± 4 and −6 ± 2 Hz, respectively). In contrast, bare TiO₂ nanoparticles did not bind to +PG membranes (+ 0.5 ± 2 Hz, Figure 3B). A discussion of the binding of bare TiO₂ NPs to PC, +PG, and +Chol membranes, as well as photocatalytic effects under UV illumination can be found in.[22] Thus, the extensive adsorption of LL-37-coated TiO₂ NPs to anionic +PG membranes can be directly attributed to the cationic peptide coating of the NPs. A possible adsorption of free peptide (i.e., not bound to the NPs surface), leading to comparable shifts in the frequency, was ruled out through a control experiment (Figure S12, Supporting Information).

After NP binding, the bilayers were exposed to UV illumination (Figure 4A,B). Mirroring the result on NP binding, only minor variations of frequency (i.e., |ΔF|<7 Hz) and dissipation (i.e., |ΔD|<1·10⁻⁶) were found for PC and +Chol bilayers after UV exposure for 2 h in the presence of LL-37-loaded TiO₂ NPs

<figure><img src="image_7.png" /><figcaption>**Figure 4.** A) QCM-d results showing the Frequency Shift (Hz) induced by LL-37-TiO₂ NPs on supported PC, +Chol, and +PG bilayers with or without UV illumination. B) Corresponding effects of bare and LL-37-TiO₂ NPs on supported +PG bilayers. All measurements were performed in 10 mM acetate, pH 5.4, in triplicate. Representative QCM-d profiles are reported in Figure S11B (Supporting Information).</figcaption></figure>
<figure><img src="image_8.png" /><figcaption>**Figure 5.** Neutron reflectivity curves with best model fit (upper) and corresponding SLD profiles (lower) for supported +PG bilayers before and after incubation with bare TiO₂ NPs (left), LL-37-coated TiO₂ NPs (middle), or for supported PC bilayers before and after incubation with LL-37-coated TiO₂ NPs (right). Shown are also the reflectivity curves with the best model fit and SLD profiles for the corresponding systems after 2 h of in situ UV exposure. All experiments were performed in 10 mM acetate buffer, pH 5.4, at a nanoparticle concentration of 100 ppm. Curves are shown for two different buffer contrasts (DAC and HAc) and the data for the latter are offset by 1·10⁻⁵ for clarity. A small offset is also applied between NR profiles in the same buffer, obtained before, after NPs addition, and after 2 h of UV illumination. The grey box in the SLD profiles indicates the position of the silicon block and reflecting interface, consisting of bulk Si and a SiO₂ layer.</figcaption></figure>

(Figure 4A; see also representative QCM-d profiles in Figure S11B, Supporting Information). In contrast, a dramatic increase in frequency (38 ± 5 Hz) was observed for +PG, indicating a massive loss of material from the surface. In parallel, significant decreases in dissipation were observed, suggesting the formation of a more diffuse adsorbed layer (Figure S11B, Supporting Information). These results indicate that LL-37-coated TiO₂ NPs induce major structural rearrangements of +PG bilayers, resulting from UV-induced oxidative degradation, but very limited structural destabilization of PC and +Chol membranes. Similar results were obtained at pH 7.4, showing that LL-37 coatings enhanced nanoparticles binding to +PG (Figure S13A, Supporting Information), as well as UV-induced oxidative degradation (Figure S13B, Supporting Information).

### 3.3. Consequences of Lipid Oxidation for Membrane Structure and Destabilization

To obtain more detailed clues to how nanoparticle-induced oxidation translates into structural changes in the lipid bilayers, NR

experiments were performed. For this, experiments were carried out at pH 5.4, using acetate buffer prepared either in H₂O (HAc) or D₂O (DAC). Results obtained under the two contrasts were fitted simultaneously to a 4-layer model (Model 1 in Scheme S1), with the input parameters reported in Tables S1 and S2 (Supporting Information). To reduce the number of free parameters, the head group thickness was fixed at 7.5 Å.[52] For all three membrane compositions, the initial bilayers presented >99% coverages and low (≈0%) tail hydration. The best-fitting parameters for the +PG and PC bilayers are reported in Table S2 (Supporting Information). Values of area per molecule (APM) and surface coverage (Γ), calculated as described previously,[22] are consistent with previous literature.[53,54]

Shown in Figure 5 are reflectivity data with best curve fits and corresponding SLD profiles for +PG bilayers interacting with 100 ppm bare TiO₂ (left panel) or LL-37-TiO₂ NPs (middle panel), and for PC bilayers interacting with LL-37-TiO₂ NPs (right panel), before and right after 2 hours of UV illumination. Reflectivity data and corresponding SLD profiles of +PG and PC bilayers before NP addition are also shown for comparison. Key structural data obtained from neutron reflectometry fits are plotted in
<figure><img src="image_9.png" /><figcaption>**Figure 6.** Structural effects on supported +PG bilayers with bare TiO₂ or LL-37-TiO₂ NPs, and on supported PC bilayers with LL-37-TiO₂ NPs. The reported parameters were extracted from neutron reflectometry fits for the three systems:[1] before nanoparticle incubation;[2] after nanoparticle incubation;[3] after 2 h in situ UV exposure. Shown are changes in A) bilayer thickness, B) area per lipid molecule (APM), C) surface coverage of the supported lipid bilayer (Γ), D) hydration of the hydrophilic headgroups, and E) hydration of the hydrophobic tails. Experiments were performed in 10 mM acetate buffer, pH 5.4, at a nanoparticle concentration of 100 ppm.</figcaption></figure>

**Figure 6** and listed in full in Table S2 (Supporting Information). Without UV illumination, no structural modifications were detected for +PG bilayer exposed to bare TiO₂ NPs (Figure 5, left panel), as evidenced by the preservation of its initial structural parameters (Figure 6). In contrast, the incubation with LL-37-TiO₂ NPs (prior UV illumination) caused significant modifications of the +PG structure (Figure 5, middle panel). Specifically, the area per molecule (APM) for +PG changed from 64 ± 3 Å² before nanoparticle addition to 173 ± 7 Å² after the addition of TiO₂ NPs coated with LL-37, which was accompanied by increased hydration of both the polar headgroups and acyl chain regions of the bilayer (Figure 6). A considerable lipid removal was associated with this, with surface coverage decreasing from 4.0 ± 0.1 mg m⁻² for the initial bilayer, to 1.5 ± 0.1 mg m⁻² after the incubation with LL-37-TiO₂ NPs. In addition, a rough NP outer layer was formed on the surface of the lipid bilayer, as reported also previously[22] and sketched in Scheme S1 (Model 2), the latter described

by the structural parameters listed in Table S3 and plotted in Figure S14 (Supporting Information). In line with QCM-d results (Figure 3B), the occurrence of this layer (absent for +PG interacting with bare TiO₂ NPs) is connected to the much higher adsorption of LL-37-coated NPs as compared to bare ones. In contrast to the large effects observed for +PG bilayers, only minor variations in the reflectivity profiles and corresponding SLDs were observed for PC (Figure 5, right panel), associated with negligible changes in the AMP and surface coverage (Figure 6, and Table S2, Supporting Information). The NPs outer layer, formed upon LL-37-TiO₂ NPs adsorption, showed slightly lower thickness and higher hydration as compared to the one formed on +PG bilayers (Figure S14, and Table S3, Supporting Information), highlighting lower adsorption of LL-37-TiO₂ NPs, in line with QCM-d data (Figure 3A).

For +PG bilayers, some structural variations were observed as induced also by bare TiO₂ NPs upon 2 hours of UV illumination,
<figure><img src="image_10.png" /><figcaption>**Figure 7.** Structural parameters, obtained from neutron reflectometry fits (Figure S15, Supporting Information), of +PG supported lipid bilayer before, and after the incubation of either bare or LL-37-coated TiO₂ NPs, as well as at different time points during exposure to UV illumination. Shown are the results of the calculated A) adsorbed amount (Γ), B) the area per molecule (APM), C) as well as of the hydration of the hydrophilic headgroups and D) hydrophobic tails.</figcaption></figure>

with an increase of the AMP from 64 ± 3 Å² to 89 ± 4 Å² (Figure 6). This corresponds to a partial bilayer removal, with the final surface coverage changing from 3.98 ± 0.04 mg m⁻² (before UV) to 2.8 ± 0.1 mg m⁻² (after UV). However, a much more pronounced destabilization of +PG was observed upon UV illumination in the presence of LL-37-TiO₂ NPs, with the hydration of the acyl chains increasing from 48 ± 4% to 88 ± 1%, paralleled by a similar increase in the polar headgroups hydration (from 74 ± 10% to 87 ± 2%). As a consequence of this, the final APM values increased to 419 ± 7 Å², corresponding to an almost quantitative bilayer removal, with a final surface coverage of 0.60 ± 0.06 mg m⁻². In contrast, LL-37-TiO₂ NPs only caused a negligible UV-induced structural destabilization of PC bilayers, leaving all key structural parameters of the lipid bilayer essentially unaffected. This demonstrates that LL-37-TiO₂ NPs show a high selectivity for +PG bilayers compared to PC in terms of UV-induced oxidative degradation, in line with the QCM-d results on the same systems (Figure 4A).

Next, reflectivity changes were monitored during UV illumination over a narrower Q-range for faster data acquisition.[55] Results for +PG interacting with bare or LL-37-TiO₂ NPs are shown in Figure S15 (Supporting Information) while corresponding data for PC are shown in Figure S16 (Supporting Information). As shown in Figure S15 (Supporting Information) (top panel), relatively small effects were observed for bare TiO₂ NPs. In contrast, pronounced decreases in reflectivity and correspond-

ing SLD variations were observed over time for LL-37-TiO₂ NPs (Figure S15, Supporting Information, bottom panel). Analysis of these results (Figure 7) showed that the destabilization of +PG induced by LL-37-TiO₂ NPs under UV illumination involved an initial incubation phase of ≈ 20 min, characterized by relatively stable values of headgroups hydration, AMP, and surface coverage, paralleled by mild variations in the acyl chains hydration. After that, a fast oxidative lipid removal, mirrored by a steep increase in the AMP, was observed on subsequent UV exposure. This process slowed down only after 100 min of UV exposure when the surface coverage value approached 0 mg m⁻². Remarkably, the acyl chains region underwent significant changes in the first 40 min of UV exposure as compared to the polar headgroups, suggesting that the oxidative destabilization induced by LL-37-TiO₂ NPs upon UV originated in the hydrophobic bilayer core. A similar analysis for PC bilayers interacting with LL-37-TiO₂ NPs is reported in Figure S17 (Supporting Information), showing no detectable variations in the bilayer structural parameters throughout 2 hours of UV exposure.

### 3.4. Antibacterial Effects and Cell Toxicity

In order to investigate if the results for model lipid bilayers translated to bacterial and human cell membranes, we next investigated antimicrobial effects of bare and peptide-coated TiO₂ NPs,
<figure><img src="image_11.png" /><figcaption>**Figure 8.** Representative confocal microscopy images obtained using LIVE/DEAD assay (red, green, and Differential Interference Contrast images overlaid) for 10⁸ CFU mL⁻¹ of *E. coli* and *S. aureus* bacteria in 10 mM acetate, pH 5.4, without or with 1 hour of incubation with bare (middle) or LL-37-coated (right) TiO₂ NPs with or without UV illumination. For each system, the fluorescence from alive (green) and dead (red) bacteria is shown in separate images (left images for alive bacteria and right images for dead ones).</figcaption></figure>

as well as their toxicity against human cells. In order to monitor antimicrobial effects, confocal microscopy was employed to study the effects of bare and LL-37-coated TiO₂ NPs on *E. coli* and *S. aureus* bacteria. For this, a two-color fluorescence LIVE/DEAD assay was used, which allows for separating the fluorescence from bacteria with intact cell membranes (staining green) from the ones with damaged membranes (staining red). Representative confocal microscopy images, separately showing intact (green-stained) and dead (red-stained) bacteria, and corresponding data, are shown in **Figures 8** and **9A**, respectively. In the absence of UV illumination, bare TiO₂ NPs had negligible effects on Gram-negative *E. coli* (3 ± 1% dead bacteria), while displaying some antimicrobial action against Gram-positive *S. aureus* (24 ± 6% dead bacteria). For both *E. coli* and *S. aureus*, the decrease in viability observed upon UV illumination was similar in the presence and absence of bare TiO₂ NPs, indicating relatively minor effects of the latter. In contrast, LL-37-coated TiO₂ NPs strongly promoted bacteria killing already prior to UV illumination, particularly so for *E. coli*, leading to a dead bacteria percentage of 68 ± 5% for *E. coli* and 54 ± 1% for *S. aureus*. UV illumination further enhanced the antimicrobial efficiency of LL-37-TiO₂ NPs, leading to 88 ± 5% and 69 ± 3% dead *E. coli* and *S. aureus*, respectively.

Possible cytotoxic effects of nanoparticles against eukaryotic cells were addressed by lactate dehydrogenase experiments em-

ploying human THP1 monocytes. For this, THP1 monocytes were exposed to bare and LL-37-coated TiO₂ NPs with or without UV illumination. Contrasting the boosting of antimicrobial effects by LL-37 coating, lactate dehydrogenase experiments demonstrated that both bare and LL-37-coated TiO₂ NPs (either before or after UV illumination) did not induce any significant increase in membrane rupture and resultant release of intracellular lactate dehydrogenase (Figure 9B). Together, these results indicate that the boosted photocatalytic destabilization by LL-37-coated TiO₂ NPs, found for bacteria-mimicking model lipid membranes, translate also to the biological systems. Furthermore, they demonstrate that AMP coating indeed provides an approach to increase the selectivity of photocatalytic TiO₂ NPs to bacterial membranes.

## 4. Discussion

Cationic surface modification of nanomaterials is a frequently employed approach for enhancing their antimicrobial effects. Since anionic phospholipids are abundant in bacterial membranes, as are negatively charged lipopolysaccharides, such surface modifications result in an increased driving force for nanoparticle binding to bacterial membranes, as well as boosted particle-mediated membrane destabilization.[³⁻⁵] At the same
<figure><img src="image_12.png" /><figcaption>**Figure 9.** A) Quantification of confocal microscopy images, showing percentages of dead *E. coli* and *S. aureus* bacteria with or without UV illumination. Results are shown both for untreated bacteria and for bacteria exposure for 1 hour to TiO₂ or LL-37-TiO₂ NPs. B) LDH release from human THP1 monocytes in the absence and presence of bare TiO₂ or LL-37-coated TiO₂ NPs, with or without UV illumination. The dashed line represents the background release in buffer. Measurements were performed in triplicate.</figcaption></figure>

time as cationization may enhance antimicrobial effects under some conditions, however, unselective binding of anionic proteins may cause corona formation,[56,57] which in turn may result in decreased antimicrobial effects. In addition, cationic surface modification frequently results in destabilization also of human cells, thereby inducing toxicity.[3,58,59] To increase selectivity of such surface modification of antimicrobial nanomaterials, it is therefore key to search for those that are able to selectively boost nanoparticle destabilization of bacterial membranes, leaving membranes of human cells unaffected. Of interest to the present study, AMPs may display potent antimicrobial effects, also against strains resistant to antibiotics.[25,28,29] It is therefore interesting to investigate to what extent such selectivity can be translated to antimicrobial nanomaterials by their surface coating by such peptides. Here, it should be remembered, however, that antimicrobial effects of AMPs rely on peptide binding and insertion into the membrane, sometimes associated with conformational changes, as well as effects on anionic lipid recruitment, domain formation, and flip-flop rates.[25] As these effects can all be expected to be partially impaired by constraining the AMP to the particle surface, it is unclear how much of the selectivity displayed by the free peptide can be transferred to the coated nanoparticles. Considering this, different approaches have been reported for increasing antimicrobial effects of surface-bound AMPs, notably by tethering/loading the

peptides to surface-bound polymer chains/matrices, thereby increasing the mobility freedom of the AMP, approaching that of the free peptide.[60,61] While little is known on how peptide orientation, packing density, mobility freedom, and conformational changes for surface-bound AMPs affect their antimicrobial effects, and the selectivity between bacteria and human cells, AMP coating of nanomaterials has been reported in a couple of studies to increase antimicrobial potency and selectivity,[62,63] as well as to provide additional features of interest, such as aggregation of bacteria and bacterial lipopolysaccharides for localization of infection and inflammation.[64] Similarly, anti-cancer effects displayed by some AMPs in solution may be transferred to nanoparticles by peptide coating for selective targeting of cancer cells.[65]

For photocatalytic nanomaterials, boosting antimicrobial effects and membrane selectivity by AMP coating is, however, more complex than for other nanomaterials. Thus, the peptide coating may result in an effectively reduced particle/water surface, and hence also reduced possibilities for ROS formation. Furthermore, the peptide may be degraded by generated ROS, and the particle coatings are effectively destroyed on illumination. Despite these concerns, a couple of studies have reported advantageous effects on coating photocatalytic nanomaterials with antimicrobial peptides and other macromolecules. Thus, Lu et al. reported dimethylpyrrolidonium-coated C₆₀ to strongly
suppress bacteria in wounds infected with *P. aeruginosa* during illumination. This, in turn, resulted in a dramatic improvement in survival, from 8% to 82% survival after treatment.[66] Similarly, Zhang et al. showed peptide-fullerene hydrogels to potently suppress multi-drug resistant *S. aureus* and promote wound healing.[67] Moreover, Chen et al. found ZnO nanoparticles coated by the AMP UBI<sub>29-41</sub> to enhance the antimicrobial effect of vancomycin against *S. aureus* and *B. subtilis*.[68] Extending on these studies, as well as on our previous studies on bare TiO<sub>2</sub> nanoparticles,[22,23] the present investigation provides a mechanical foundation for observations of retained or improved antimicrobial effects of photocatalytic nanoparticles after AMP coating. In particular, it shows that peptide coating may be done without substantially suppressing or altering ROS generation on illumination. While the ζ-potential results indicate that the peptide layer on TiO<sub>2</sub> NPs undergoes some oxidation on UV exposure, which may include surface-bound peptide participating in the reaction through the generation of reactive oxygen or nitrogen species, particle size and ζ-potential results also show peptide-coated dispersions to be colloidally stable, and the layers thus stable with respect to peptide desorption over the duration of the experiments (≈hours). Hence, there is no experimental support for the peptide layers effectively desorbing to catalyze oxidative degradation of either model lipid or bacteria/cell membranes. Instead, the QCM-d results clearly show that the main effect of the peptide coating (remaining positively charged also after UV exposure) is to promote binding to bilayers enriched in anionic PG. Due to the short lifetime of ROS, close proximity between ROS generated and their lipid target molecules affords AMP-coated TiO<sub>2</sub> nanoparticles with promoted capacity to destabilize lipid membranes under UV illumination. Apart from outlining the details of such oxidative destabilization of lipid membranes in detail for model lipid systems, the antimicrobial and cell toxicity results provided in this investigation demonstrate such effects to translate also to the biological situation.

However, there are still numerous issues that will have to be resolved regarding the selectivity of peptide-coated photocatalytic nanoparticles. For example, qualitatively similar effects were observed for the AMP LL-37 and for poly(arginine) homopolypeptide (Figure S18, Supporting Information), but it remains to be investigated how the different antimicrobial potency and selectivity displayed by these as free peptides in solution[69–71] translate into more detailed biological performance. Furthermore, the antimicrobial action of LL-37 has been correlated to the formation of an amphiphilic helix conformation,[69] an effect likely to be (partly) lost after peptide immobilization at the nanoparticle surface. Considering this, membrane interactions may be governed by the distribution of charges, as well as hydrophilic and hydrophobic amino acids, at the surface of peptide-coated nanoparticles. Addressing these aspects is therefore a key issue in further studies.

In addition, bacterial lipopolysaccharides play a key role not only in the structural integrity of bacteria but also in their pathogenicity. Investigating the degradation of lipopolysaccharides by bare as well as peptide-coated nanoparticles is therefore key for a complete understanding of photocatalytic antimicrobial effects of such systems. So far, only a couple of studies have been reported, suggesting that lipopolysaccharides provide further complexity to photocatalytic antimicrobial effects of

nanoparticles by: i) having a lower susceptibility to photocatalytic degradation than phospholipids, and ii) providing a barrier affecting the proximity between nanoparticles and the plasma membrane of bacteria.[9] In this context, AMPs again offer interesting opportunities since some of these have been found to be able to bind bacterial lipopolysaccharides and suppress their inflammatory properties,[26,72,73] a key feature for suppressing inflammation following the generation of bacterial debris after membrane lysis by antimicrobial peptides.

From a wider perspective, we note that the approach taken in the present study to boost the antimicrobial effect by promoting proximity between bacteria and antimicrobial nanoparticles may also have a bearing also for other pathogens, other types of nanoparticles, and even other biological questions. For example, photocatalytic nanoparticles have been found to display potent antiviral effects on illumination,[8,9,74,75] hence surface modifying these with compounds having an affinity for virus capsids or capsid envelopes, such as aptamers, antiviral peptides, or antibodies toward capsid proteins may offer a way to boost antiviral effects of photocatalytic nanoparticles without simultaneously increasing cell toxicity. Indeed, this is supported by findings by Hu et al., who immobilized aptamers to GO and found this to promote virus binding and result in boosted inactivation of bacteriophage MS2 via oxidative damage under illumination.[76] Analogously, one could imagine that bacteria targeting can promote antimicrobial effects also for non-photocatalytic nanoparticles.[3] Exemplifying this for nanoparticles causing membrane disruption by particle binding, Malekkhaiat Häffner et al. coated spiky, virus-like, mesoporous silica nanoparticles with the antimicrobial peptide LL-37 and found peptide coating to strongly boost membrane disruption and antimicrobial effects caused by such spiky nanoparticles,[35] analogous to edge effects caused by sheet nanoparticles.[77] Similarly, one could envision nanoparticles reaching antimicrobial effects through local heating induced by either light or magnetic field[3] to benefit from enhanced proximity between bacteria and nanoparticles through targeting surface coatings, e.g., based on antimicrobial peptides.[78,63] Finally, we note that photocatalytic and other nanomaterials are interesting not only for combatting infection but also in other biological contexts, such as in tumor therapeutics. Exemplifying this, Duong et al. demonstrated doxorubicin-containing polymer nanoparticles coated with the antimicrobial peptide GRR10W4 to display improved uptake and toxic effects for melanoma cancer cells, but not for non-malignant fibroblasts and keratinocytes.[65] Considering this, as well as the promising results in the present study, investigations into these lines of inquiry seem relevant.

From a mechanistic perspective, key developments in nanoparticle-membrane interactions for other types of nanoparticles may be of relevance also for photocatalytic NPs, either bare or coated.[9,79,80] For example, inspiration can be found in plasmonic gold nanoparticles and how to investigate the dynamics and exchange of capping ligands for such nanoparticles, nanoparticle size effects, as well as how these influence membrane interactions and antimicrobial effects.[79–82] Furthermore, preferential nanoparticle interactions with membrane domains[83,84] are likely important for photocatalytic NPs and deserve further attention, as do the effects of nanoparticles on integrin activation and other key membrane processes by systematically varied nanoparticle properties.[85] From this perspective,
<figure><img src="image_13.png" /><figcaption>**Figure 10.** Schematic illustration of experimental results demonstrating the feasibility of coating photocatalytic NPs with AMPs for selective boosting of antimicrobial effects: On coating TiO₂ NPs by positively charged LL-37 at pH 5.4-7.4, the peptide coating does not detrimentally interfere with ROS generation and displays good stability on UV exposure for 1–2 h. As a result of this, binding of peptide-coated TiO₂ is much higher than that of bare TiO₂ NPs, particularly at anionic bacteria-like membranes, and oxidative degradation of the latter on UV exposure strongly boosted by peptide coating, whereas corresponding effects on zwitterionic mammalian-like membranes were much smaller. Mirroring this, LL-37-coated TiO₂ NPs displayed boosted antimicrobial effects against both Gram-negative *E. coli* and Gram-positive *S. aureus* bacteria, whereas toxicity against human THP1 monocytes remained low.</figcaption></figure>
we note that the platform described in the present investigation
is compatible with all such future research directions.

**5. Conclusions**

As indicated by ζ-potential measurements, TiO₂-bound LL-37 remains partially intact even after UV illumination for 1–2 hours. Furthermore, ROS generation on illumination of TiO₂ NPs is not detrimentally affected by AMP coating, as assessed from C₁₁-BODIPY fluorescence. QCM-d showed that coating with positively charged peptides promotes the binding of TiO₂ NPs to anionic bacteria-mimicking membranes, thereby boosting oxidative membrane degradation during UV illumination. In contrast, zwitterionic mammalian-like membranes displayed much lower binding of LL-37-coated TiO₂ NPs and were much less susceptible to oxidative degradation. NR results showed such membrane destabilization to involve increased hydration, lipid removal, and solubilization. Mirroring these physicochemical results obtained for model lipid membranes, antimicrobial effects under UV illumination were found to be promoted after LL-37 coating for both E. coli and S. aureus. Meanwhile, toxicity against human THP1 monocytes remained low for the LL-37-coated TiO₂ NPs. Together, these findings illustrate that AMP coating offers opportunities for selectively boosting the antimicrobial effects of photocatalytic nanoparticles at retained low toxicity (Figure 10).

**Supporting Information**

Supporting Information is available from the Wiley Online Library or from
the author.

**Acknowledgements**

The authors thank ILL (Proposal number 9-13-924) and ISIS (https://doi.org/10.5286/ISIS.E.RB2210096) for access to neutron beam time and solid-liquid interface cells. The research was funded by the Swedish Research Council (Grant numbers 2021-05498; MM, LC), Independent Research Fund Denmark (Grant number 9040-00020B; MM and EPO), as well as the LEO Foundation Center for Cutaneous Drug Delivery (Grant number 15007; MP and MM). The Swedish Research Council and the Swedish Foundation for Strategic Research were also acknowledged for access to nCHREM via ARTEMI, the Swedish National Infrastructure in Advanced Electron Microscopy (Grants No. 2021-00171 and No. RIF21-0026). Funding Sources: Neutron beam time was awarded at ILL (Proposal number 9-13-924) and ISIS (RB2210096). The research was funded by the Swedish Research Council (Grant numbers 2021-05498; MM, LC), the Independent Research Fund Denmark (Grant number 9040-00020B; MM, EPO, and LC), as well as the LEO Foundation Center for Cutaneous Drug Delivery (Grant number 15007; MP, MM).

**Conflict of Interest**

The authors declare no conflict of interest.

**Author Contributions**

L.C. and E.P.O. contributed equally to this work. E.P.O., L.C., and M.M. designed the study. All authors were involved in performing the experiments and the interpretation of the experimental results. L.C. and M.M. wrote the paper. All authors reviewed the paper.

**Data Availability Statement**

The data that support the findings of this study are available in the supplementary material of this article.

**Keywords**

Antimicrobial peptides, lipid membranes, lipid oxidation, LL-37, photocatalysis, TiO₂

Received: October 19, 2023

Revised: January 30, 2024

Published online: February 25, 2024

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