# Unraveling the complexity of surface antibacterial effects: A multifaceted evaluation of electrodeposited nanospikes

Marion Stalet, Sara Hélis, Laurence Convert, Rémi Dreyfus, Jean-François Bryche, Loïc Leroy, Paul Charette, Michael Canva, Yoann Roupioz

► To cite this version:

Marion Stalet, Sara Hélis, Laurence Convert, Rémi Dreyfus, Jean-François Bryche, et al. Unraveling the complexity of surface antibacterial effects: A multifaceted evaluation of electrodeposited nanospikes. Applied Surface Sciences Advances, 2025, 30, pp.100881. 10.1016/j.apsadv.2025.100881. hal-05349461

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Submitted on 5 Nov 2025

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# Unraveling the Complexity of Surface Antibacterial Effects: A Multifaceted Evaluation of Electrodeposited Nanospikes

Marion Stalet<fnref n="1" />,<fnref n="2" />,<fnref n="3" />, Sara Hélis<fnref n="1" />,<fnref n="2" />, Laurence Convert<fnref n="1" />,<fnref n="2" />, Rémi Dreyfus<fnref n="1" />,<fnref n="2" />, Jean-François Bryche<fnref n="1" />,<fnref n="2" />, Loïc Leroy<fnref n="3" />, Paul G. Charette<fnref n="1" />,<fnref n="2" />, Michael Canva<fnref n="1" />,<fnref n="2" />, Yoann Roupioz<fnref n="3" />

Corresponding author: yoann.roupioz@cea.fr

Additional corresponding authors: marion.stalet@usherbrooke.ca, michael.canva@usherbrooke.ca.

## Abstract

The intricate interaction between bacteria and surfaces, coupled with the complexity of biofilm formation, poses significant challenges in understanding and controlling microbial behaviour. To combat pathogen contamination and proliferation, bioinspired nanostructures as antimicrobial surfaces have gained prominence over the past decade. However, conflicting theoretical studies and experimental results have made it difficult to draw general conclusions about antibacterial mechanisms. To address these challenges, we have adopted an approach that provides access to deeper insights to characterize the performance of surfaces, by taking into account different modes of antibacterial action. To demonstrate the applicability of this strategy, nanostructured materials mimicking dragonfly wing, were evaluated. A gold electrodeposition process was optimized to create large-area, uniform nanospike arrays, ranging in height from 70 to 570 nm. Large sample area (12 cm<fnref n="2" />) was critical for ensuring statistically significant results in the analysis of antibacterial effects. Model surfaces with ~150 nm high spikes were tested, demonstrating significant efficacy in reducing bacterial proliferation for *Escherichia coli* and *Staphylococcus epidermidis*. Moreover, striking differences between strains were observed in biofouling (surface detachment and release of bacteria). These findings underscore the relevance of such a protocol in precisely characterizing bacterial interactions with antimicrobial materials. By providing a standardized characterization method, this study aims to facilitate a deeper understanding of antimicrobial surface interactions with bacteria and contributes to the development of more effective antibacterial surfaces.

## Keywords

Antibacterial surfaces; Cell-surface interactions; Nanostructuring process; Electrodeposition; Mechano-bactericidal.

<footnote n="1">¹Laboratoire Nanotechnologies Nanosystèmes (LN2 – IRL 3463), CNRS, Université de Sherbrooke, Sherbrooke, QC J1K OA5, Canada</footnote>

<footnote n="2">²Institut Interdisciplinaire d'Innovation Technologique (3IT), Sherbrooke, QC J1K OA5, Canada</footnote>

<footnote n="3">³Univ. Grenoble Alpes, CEA, CNRS, IRIG-SyMMES, 38000 Grenoble, France.</footnote>
# 1. Introduction

Surface contamination and biofilm formation pose significant challenges in many areas, including food safety, water and air treatment, public transportation, cooling systems, and many others. These issues are particularly critical in healthcare, where the proliferation of bacterial pathogens on medical devices and implants can lead to serious patient infections.[1] Biofilms are structured communities of microorganisms embedded in a self-produced extracellular polymeric matrix, which provides advantages such as enhanced protection and intercellular communication. Their formation is a highly complex and dynamic biological process that is not yet fully understood, as it involves a series of stages such as initial adhesion, microcolony formation, maturation, and eventual detachment or dispersal of microorganisms.[2,3] Once established, biofilms exhibit strong resistance to antibiotics and conventional treatments, making early intervention crucial to prevent persistent contamination and infections. The rising prevalence of antibiotic resistance, coupled with the wide variety of pathogens capable of contaminating surfaces, underscores the urgent need for innovative antimicrobial strategies. Ideally, these solutions should be effective against a broad spectrum of microorganisms while remaining safe for humans and the environment. Moreover, given their widespread applications, these solutions should be large-scale, cost-effective and require minimal maintenance to ensure long-term efficacy.[4]

Over the past decade, micro and nanostructured surfaces emerged as robust solutions to address microbial contamination challenges, drawing inspiration from natural active surfaces.[5] Initially investigated by Ivanova et al. in 2012, bioinspired surfaces showed mechano-bactericidal effects, reducing bacterial contamination passively, without releasing antimicrobial molecules into the environment to which bacteria could eventually develop resistance, and efficiently targeting several species of microorganisms.[6] Thanks to this pioneering work, it has been shown that the efficacy of such surfaces is closely related to the nature of its material and its topological characteristics.[5,6] Numerous materials, fabrication processes, and geometric parameters such as aspect ratio, shape, height, and density of nanostructures have been investigated to understand their impact on antimicrobial efficacy.[7]

Biomaterial interactions with living organisms, particularly bacteria, are inherently complex and can be deeply influenced by experimental factors, such as incubation conditions, viability assessment methods, bacterial morphology and physiological state. In addition, antimicrobial activity of surface may involve different mechanisms that must be properly evaluated. For instance, anti-adhesive surfaces prevent bacterial attachment, while self-cleaning coatings enable the easy removal of contaminating particles such as dust and bacteria, as the low adhesion of water droplets causes them to roll off the surface.[6] Bacteriostatic treatments inhibit bacterial growth and reproduction without killing, allowing for potential bacteria recovery after detachment. In contrast, contact-killing surfaces are bactericidal only upon direct contact, whereas surfaces with drug-release mechanisms either kill or inhibit bacteria by continuously releasing antimicrobial compounds. This latter approach is thus time dependent and controlled by the amount of drug loaded (and released) from the material surface. In many cases, the mode of action is closely intertwined with environmental conditions: for example, metal-ion release coatings or drug-release surfaces require the presence of moisture to activate and sustain their antimicrobial effect. Recognizing these dependencies is critical, as variations in hydration or incubation conditions can strongly alter performance. Furthermore, many antimicrobial activities
studies, including some established ISO and ASTM methods discussed in section 3.2.1, rely on a
single characterization method, such as fluorescence imaging or colony forming unit (CFU)
counting in liquid media, which provides only a partial assessment of the overall antimicrobial
properties of the surface. As a result, experimental data published so far are often very specific to
test conditions, making it difficult to extrapolate results to other surface designs or different
pathogen panels, and may even be contradictory.[8,9] At the same time, specificity is necessary to
match the intended application of these materials, or to test under worst-case scenarios, to ensure
that results remain clinically or industrially relevant. However, despite the large number of
studies evaluating the antimicrobial activity of nanostructured surfaces, the mechanisms
underlying the mechano-bactericidal mode of action are not yet fully understood. In order to
develop highly effective antimicrobial surfaces and deepen our understanding of their interactions
with microorganisms, it is essential to adopt standardized characterization protocols. They must
account for the multiple antibacterial mechanisms at play and minimize experimental bias to
allow meaningful comparisons between studies.

**A ELECTRODEPOSITION NANOSTRUCTURING**

<figure><img src="image_2.png" /></figure>

**B MICROBIOLOGICAL CHARATERIZATION**

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

**C ANTIBACTERIAL SURFACE PROPERTIES**

<figure><img src="image_4.png" /><figcaption>Figure 1 – Schematic overview of the fast electrochemical nanostructuration and anti-microbial activity assessment; A. Electrodeposition parameters optimization to obtain dragonfly bioinspired nanostructuration. B. General protocol for antimicrobial properties characterization. The bacteria inoculum is deposited onto the sample surface containing nanostructured zones and smooth gold as the reference. Step 1: adhesion; Step 2: reproliferation. C. Visual representation of the different properties under the “antibacterial” umbrella.</figcaption></figure>

In this study, the objectives are to address these challenges by developing a comprehensive
methodology for the full characterization of antibacterial surface properties and demonstrating its
effectiveness on gold nanospikes, as a proof of concept. We first fabricated 12 cm² nanostructured
surfaces using a gold electrodeposition process previously used for microsensors, optimizing it
for uniformity and controlled nanostructure formation over centimetric areas.[10][11] These
structures were designed to mimic the antimicrobial nanospikes found on dragonfly wings and
typically ranged from 150 to 200 nm in height (Figure 1A). The properties of the antimicrobial
engineered surfaces were evaluated against common biomedically relevant pathogens,
specifically *Escherichia coli* and *Staphylococcus epidermidis* (see Figure 1B). In this work, the
term antimicrobial was used in a broad sense, as the protocol could in principle be extended beyond bacteria to other microorganisms. While the present study focuses on bacteria, some adaptations could allow testing against filamentous fungi or even viruses. Rather than using a one-step labeling of nuclei and membrane to assess bacterial membrane integrity upon antimicrobial surface interaction, we applied an in-depth characterization protocol that evaluated several key physiological effects such as bacterial initial adhesion, proliferation in medium and on the surface, biofilm formation and even re-proliferation after adhesion to the surface. These information were easily obtained by completing viability assays on bacteria adhering on surfaces and by evaluating the viability of planktonic cells interacting with the nanostructured surfaces. An ultrasound-free protocol based on a combination of adhesion and proliferation assays was designed, as widespread methods such as ultrasound detachment have been shown to exhibit repeatability and reproducibility issues and affect bacterial viability.[12,13] A first incubation step in a poor and bacteriostatic medium was used to evaluate the capacity of planktonic bacteria to adhere to surfaces (step 1). By combining fluorescent labeling with solid agar enumeration, both the density of surface-bound and unbound bacteria counts were characterized in parallel. After this first characterization, the incubation of inoculated surfaces in a rich culture medium was carried as a reproliferation assay to monitor changes in bacterial viability and concentration along with their ability to detach from the surfaces (step 2).

Through this investigation, we proposed a comprehensive characterization protocol as a reference method for assessing the various antibacterial properties of nanostructured surfaces, as displayed in *Figure 1C*, using promising electrodeposited surfaces to combat surface contamination by bacterial pathogens. While this study focused on bacteria, the approach could be easily adapted to investigate other microorganisms such fungi, algae and viruses, providing a versatile framework to advance the design of active surfaces. Our goal is to deepen our understanding of the complex interactions involved in cell-surface interactions to facilitate the development of advanced antimicrobial surfaces.

## 2. Materials and methods

### 2.1 Chemicals and Reagents

2-Propanone (acetone), ethanol (≥95%), propan-2-ol (IPA), phosphate-buffered saline (PBS, pH 7.2-7.6, one tablet/200 mL), and tryptic soy broth (TSB) were purchased from Sigma Aldrich (Saint Quentin Fallavier, France). OptiClear™ cleaning solvent was obtained from National Diagnostic (Atlanta, GA, USA). Hydrogen tetrachloroaurate(III) trihydrate (HAuCl₄), lead(II) acetate trihydrate (Pb(CH₃COO)₂), and Invitrogen LIVE/DEAD™ BacLight™ bacterial viability kit were purchased from ThermoFisher (Waltham, MA, USA). Trycase soy agar (TSA) was purchased from Biomérieux (Lyon, France). All solutions were prepared using Milli-Q (18 MΩ/cm) deionized water and sterilized by autoclave for liquids before use. *Escherichia coli* (ATCC11775) and *Staphylococcus epidermidis* (ATCC12228) were purchased from the American Type Culture Collection (Manassas, VI, USA), as representative Gram-negative and Gram-positive microorganisms, respectively.

### 2.2. Nanostructured surface fabrication by electrodeposition.

#### 2.2.1. Electrodeposition of nanostructures and electrochemical characterization
Before and after each step of the fabrication, 75 x 25 mm microscope glass slides from Fischerfinest (Fisherbrand, Hampton, NH, USA) were thoroughly washed through four consecutive 5-minutes ultrasound baths in OptiClear™, acetone, IPA, and finally deionized water. After washing, substrates were deposited via e-beam evaporation with the Nanochrome™ I from Intlvac (Georgetown, ON, Canada). A three-nanometer titanium (Ti) adhesion layer was deposited at a rate of 0.05 nm/s, followed by a 30 nm gold (Au) layer at 0.2 nm/s, both applied solely to one side of the 19 cm² surface.

Gold nanospikes were fabricated using an adapted electrodeposition (ED) process from Sabri et al. [10,11,14] After cleaning and washing, slides were inserted into the electrochemical cell and used as the working electrode (WE) in a three-electrode adaptable set-up, allowing reproducible positioning of the substrates (see Figure S1, Supplementary Data). A silver-silver chloride electrode (Ag/AgCl, 3M KCl from BASi®, West Lafayette, IN, USA) was used as the reference electrode (RE) and a 12-inch graphite rod of 6 mm diameter from Northern Materials Innovations Ltd (Edmonton, AL, Canada) was used as the counter electrode (CE). About 12 cm² of the slide deposited surface were immersed into the electrolyte aqueous solution, composed of HAuCl₄ and Pb(CH₃COO)₂ salts. The Pb²⁺ ions in the electrolyte solution act as directional growth agents: their lower affinity towards the {111} gold facets orient the growth of the gold nanostructures by the reduction of Au³⁺ along these facets.[14] The gold electrodeposition was performed at room temperature and various HAuCl₄ solutions were tested, with concentrations of 6.8, 12.5, 25, 50 and 100 mM. Prior to any ED, the solutions were degassed with nitrogen (N₂) using an inlet tube for 10 minutes, and during ED, the N₂ flow was maintained over the solution. A 5-minute equilibration time was ensured under stirring. Grown gold nanostructures were electrodeposited on microscope slides using the CHI604C potentiostat from CH Instruments, Inc (Austin, TX, USA). ED parameters were set and controlled using the CH604c Electrochemical Analyzer user interface (v14.05) with the chrono-amperometry mode. The ED was performed at a continuous 0.05 V, with a Pb(CH₃COO)₂ concentration of 0.5 mM. Various deposition durations were investigated, with mainly 100 and 300 seconds being used. The current during ED was recorded in real time at a sensitivity of 0.1 A/V. After the ED, the sample was washed using the aforementioned procedure (OptiClear™, acetone, IPA, deionized water).

Cyclic Voltammetry (CV) was performed to measure the electrochemically active surface before and after each ED.[15] A platinum CE (from BASi®) was used. Each time, the three-electrode cell was dipped in 0.5 M H₂SO₄ solution under agitation at 600 rpm and three cycles from 0 to 1.6 V at a speed of 0.1 V/s with a sensitivity scale of 10 mA/V were recorded. The electro-active surface was calculated by integrating the oxidation peak of the CV and using the charge density of a monolayer of gold oxide. After CV, the samples were rinsed thoroughly with deionized water.

### 2.2.2. Morphological characterization of gold nanostructures

The nanostructure morphologies were imaged using scanning electron microscopy (SEM – ThermoFisher, Axia™ ChemiSem) and atomic force microscopy (AFM – Bruker, Dimension Icon ScanAsyst). SEM micrographs were acquired at tilts of 0° and 45°. The working distance was 8 mm or less, with a magnification of x 15000, an acquisition time of 1 µs, an intensity of 12 pA and a voltage of 30 kV. To assess the homogeneity of the nanostructured surface, at least five micrographs at different distances from the sample center were taken (see Figure S2,
*Supplementary Data*). Energy-dispersive X-ray spectroscopy (EDX) measurements were also taken using the SEM microscope.

The AFM images were acquired using the Dimension Icon Scanasyst (Veeco, Bruker) in Tapping mode with ScanAsyst-Air probes. The scanning rate was set to 0.5 Hz and at least three images per sample of 25 µm² (512 x 512 lines) were acquired. The average nanostructures height, the root mean square of the roughness (RMS) and the density of the nanostructures were measured and averaged from at least three 25 µm² large areas AFM and SEM images, using Gwyddion and Fiji ImageJ software. The mean height was calculated by setting the baseline of measurement at the lowest point of the sample, thus not considering the accumulated matter under the spikes.

For AFM analysis, three images were acquired per sample, for multiple samples, from different manufacturing batches. This sampling strategy was designed to capture both intra-sample variability (across different scanned areas), inter-sample variability (between distinct substrates), and inter-batch variability, thus providing a representative overview of the reproducibility and consistency of the nanostructuring process, while accounting for the inherent limitations of AFM in terms of scan area and acquisition time. A summary table presenting the variability observed across multiple deposition batches and experimental conditions is provided in the Supplementary Data (see Table S6), offering a broader overview of the reproducibility and robustness of the fabrication process.

## 2.3. Antibacterial activity assessment of the electrodeposited surfaces

### 2.3.1. Sample preparation

Twenty-four hours prior to the viability experiment, nanostructured samples were cleaned using an oxygen (O₂) and argon (Ar) plasma cleaner (Femto 5RF, Diener Electronic, Ebhausen, Germany) for three minutes at a pressure of 0.6 mbar, with 75% O₂ and 25% Ar. The samples were then stored in a clean environment, at room temperature. Before incubation with bacteria, the samples were sterilized by immersion in 70% ethanol for 15 minutes at room temperature and left to dry under sterile flow during one hour.

### 2.3.2. Bacterial strains and growth conditions

*Escherichia coli* strain ATCC 11775 and *Staphylococcus epidermidis* strain ATCC 12228 were cultivated using standard procedures. Bacteria were first grown on Trypticase Soy Agar (TSA) plates overnight at 37°C (streaking) and then one colony was selected and cultured overnight in 5 mL of Trypticase Soy Broth (TSB). Incubation proceeded overnight at 37 °C without agitation. To minimize potential effects of growth kinetics or cell death, this pre-culture was followed by enrichment, and the supernatant was removed before resuspending the bacteria in PBS.

To study surface activity under a “worst-case” scenario, bacterial cultures needed to be in the exponential growth phase, when bacteria are metabolically most active. Therefore, 300 µL of the overnight preculture were diluted into 9 mL of fresh TSB and incubated 2 hours for *Staphylococcus epidermidis* and 1 hour for *Escherichia coli* at 37 °C and 150 rpm.

2 mL of the bacterial inoculum were transferred into a sterile Eppendorf microtube and centrifugated 10 min at 5,000 x g (9,000 rpm) to pellet the bacteria. The supernatant was then removed and replaced by 2 mL PBS one time. The bacteria were resuspended by vortexing for a
few seconds and the bacterial inoculum was then adjusted to $OD_{600} = 0.33$ for *Staphylococcus epidermidis* and $OD_{600} = 0.49$ for *Escherichia coli* in PBS, which corresponds to approximately $2 \times 10^7$ Colony Forming Unit (CFU) per mL and $2 \times 10^8$ CFU/mL, respectively. The bacterial inocula were finally serially diluted to the tenth to try to reach the theoretical concentration of $2 \times 10^5$ CFU/mL, which was selected to avoid bacterial cluster suspensions. Inocula were plated to determine their actual concentrations by SPA.

### 2.3.3. Spot Plating Assay (SPA)

The initial inoculum concentration was confirmed by spot-plating assay.[16] Prior to the CFU spot plating, TSA plates were dried at room temperature under a sterile environment for five minutes (or 15 minutes for fresh TSA plates). Then, 200 µL out of the 300 µL of liquid media in contact with the surface were sampled. The bacterial solutions to be spotted were diluted to the tenth in fresh liquid culture medium. Four series of five two-fold dilutions were deposited on a 120 x 120 mm square Petri dish by spotting 30 µL per spot/dilution/series (120 µL per dilution in total) with a multichannel pipette. After spotting, plates were left on the bench for 15 minutes to ensure the spotted solutions absorption in the solid agar medium before being incubated. CFU counting were performed after 24 hours of incubation at 37 °C for *Staphylococcus epidermidis* and after 12 hours of incubation for *Escherichia coli*, and converted to a number of bacteria per well to facilitate comparison with surface-adhered bacteria counting.

### 2.3.4. Microbiological characterization

Before the addition of the bacterial solution to the nanostructured sample, a self-adhesive silicone chamber (from Ibidi GmbH, Gräfelfing, Germany) was placed onto the nanostructured slides. Each chamber is composed of 8 wells with dimensions $12 \times 7.75 \times 8.0 \text{ mm}^3$ ($w \times l \times h$), allowing for the replication of the microbiological characterization on separate areas of a single nanostructured surface. This device allows for the replication of the microbiological characterization on separate areas of a single nanostructured surface. Then, 300 µL of bacterial solution in PBS (either *Escherichia coli* or *Staphylococcus epidermidis*) were added to the wells and incubated at 37 °C and 100 rpm. For each nanostructure motif, duplicates were processed for the microbiological studies. To ensure comparability, both substrates were fabricated simultaneously under identical electrodeposition conditions.

After 2 hours of incubation, the two slides were treated differently according to the characterization process. One slide was used to study the anti-fouling effect and antibacterial agent release, while the other slide was used to characterize contact-killing/bacteriostatic effects and bacteria release from the surface. This second analysis was possible by incubating the slides during the first step in a poor and bacteriostatic medium (PBS) enabling bacteria survival but impeding their division, and then inducing the potential reproliferation by incubating slides in a culture medium (TSB) favoring bacterial growth (rich medium).

From the first slide, liquid phase (containing planktonic cells) were sampled, serially diluted, and plated on TSA for CFU assessment using a spot plating assay. After removing the liquid phase, the surfaces were gently rinsed with PBS and stained with the Live/Dead Baclight Kit™ (Invitrogen) according to the manufacturer's protocol. Bacteria were visualized with a fluorescence microscope (Nikon Ci-L Upright) at 20× magnification, using two filter sets, Cy3-4040C and GFP-1828A, to image dead and live bacteria, respectively. Fluorescent bacteria counting was determined from 15 photographs (5 images recorded for each well from 3 distinct
wells per assay) and expressed as the number of bacteria per well for each nanostructured slide. To determine if bacteria were able to recover from exposure to nanostructures, the second slide was first rinsed with PBS and then incubated in TSB (300 µL/well) for an additional 4 hours of incubation at 37 °C and 100 rpm to favor bacterial division and growth. After incubation, bacterial counts were proceeded in the same way as for the first step : culture media aliquots were spotted on agar plates to assess the number of viable planktonic bacteria, by SPA, in each well, and surfaces were stained with the Live/Dead kit to assess the viability and number of bacteria remaining on each surface. Overall, the characterization process lasts approximately 24 hours.

### 2.3.5. Fluorescence microscopic images analysis

To accelerate and automate the counting of bacteria using fluorescence images, a custom Python program was developed. This program facilitates semi-automatic brightness and contrast adjustments for certain images and automates the detection and counting of bacteria. Additionally, the program categorizes bacteria into three groups: those labeled solely by SYTO® 9, solely by PI, and by both dyes. Usually, cells stained by SYTO® 9 are considered viable, while those stained with PI are deemed non-viable. However, some cells may be labeled with both dyes simultaneously, and studies have shown that PI labeling of damaged membrane does not always correlate with bacterial death. To avoid double counting, bacteria expressing both fluorescent dyes are detected, subtracted from the "Dead" and "Alive" counts, and categorized as “Damaged” by the program.

## 3. Results and Discussion

### 3.1. Nanostructured surface fabrication by electrodeposition of gold

To produce relevant nanostructured materials for both characterization and practical applications, fabrication processes that can be scaled up to several square centimeters or more are essential. Commonly used materials for nanostructured surfaces – such as silicon, titanium, and polymers – are often processed using top-down techniques like lithography and etching.[17] While these methods provide high resolution and organized patterns, they typically require expensive infrastructure, long processing times, and trained personnel, making them less suitable for cost-effective large-scale applications. Other top-down methods have tried to overcome these limitations, such as nanoimprint lithography. Recent studies have demonstrated that resin-based nanostructured substrates can be engineered to exhibit both antibacterial and bactericidal properties when combined with thin functional coatings. For instance, resin nanopillar arrays coated with silicon dioxide (SiO₂) have shown a strong reduction in bacterial adhesion and viability, even without the use of chemical agents or antibiotics.[18] These nanostructured resins are well-suited for scalable low-cost fabrication, as they can be produced by nanoimprinting or molding techniques using UV-curable materials.[19] However, their use remains limited by the restricted functional range of polymers and the need for additional coating steps.

To address these limitations, several bottom-up approaches have emerged, including self-assembly of block copolymers (which still require an etching step), sol-gel processes and electrodeposition. Electrodeposition is an electrochemical process in which metal ions in solution are reduced and deposited onto a conductive substrate under the application of an electric current. Electrodeposition offers a versatile, cost-effective, and scalable alternative for fabricating
nanostructured surfaces. This bottom-up method enables the one-step creation of metallic or
composite nanostructures over large surface areas with controlled morphology and thickness,
even on non-planar substrates.[20,21] It is particularly suited for high-throughput production and
reproducibility, making it attractive for practical antimicrobial applications.[22–24] Furthermore,
electrodeposition is compatible with a wide range of materials, including noble metals (gold,
silver, platinum) and base metals (zinc, nickel, copper, etc.) as well as their oxides and
alloys.[25–27] Limitations associated with certain metals, such as reactivity or solubility, can
often be addressed using ionic liquids or non-aqueous electrolytes.[28] Moreover, surfaces
fabricated via electrodeposition exhibit inherent variability in the dimensions and shapes of the
deposited structures. This random nanostructuration contrasts with the literature focusing on well-
defined nanostructures arrays. While these investigations are crucial for elucidating key
antimicrobial parameters related to nanostructures geometry, the potential of random and
heterogeneous structures, such as nanoneedles on dragonfly wings, remains of critical interest, as
the dimension variations between the features could help achieve broad-spectrum
efficiency.[29,30]

For our study, we selected gold electrodeposition as the fabrication method due to its excellent reproducibility, ease of parameter tuning, and compatibility with biological systems. Gold presents a number of advantages for antimicrobial applications: it is chemically inert and suitable for surface functionalization. Unlike silver, which exhibit cytotoxicity toward eukaryotic cells, gold is non-toxic, making it ideal for mechano-bactericidal studies.[31] Though gold is more expensive, only nanometric quantities are needed, minimizing material costs. Additionally, its optical (plasmonic) properties offer potential for characterization and biosensing applications[32–34]

Elbourne *et al.* successfully demonstrated the potential of electrodeposition of gold on 16 mm² surface by showing that sharp gold nanostructures with heights ranging from 100 to 555 nm can significantly reduce the percentage of viable cells of Pseudomonas aeruginosa and Staphylococcus aureus by 60 to 90% after 18 hours of incubation.[11] Wu *et al.* were also able to build circular pillar-shaped gold nanostructures, rings, and nuggets by using a combination of electrodeposition and alumina templating.[35] Using fluorescence labeling, these types of nanostructures seemed to display a >99% killing efficiency on adhered S. aureus for surfaces of 1 cm². These studies highlight the antimicrobial potential of gold nanostructures fabricated by electrodeposition.

The gold electrodeposition process was performed using HAuCl₄ salts and Pb(CH₃COO)₂ as a directional agent to produce well-defined nanostructures over 12 cm² glass substrates (standard microscope slides). We investigated key deposition parameters – including gold salt concentration, deposition duration, and electrolyte agitation – to ensure uniform nanospike formation across the entire surface. The resulting structures serve as a promising platform for assessing mechano-bactericidal activity.

**3.1.1. Impact of electrodeposition parameters on nanostructures homogeneity and morphologies**

This section explores three intricately linked parameters of electrodeposition: electrolyte agitation, HAuCl₄ concentration, and deposition duration. Although not exhaustive, optimizing
the combination of these factors has been key to achieving uniform nanostructuring on our samples. Gold-covered microscope slides were electrodeposited at continuous 0.05 V with a concentration of $[Pb(CH_3COO)_2]$ of 0.5 mM. For each parameter, a range of values was tested: electrolyte agitation from 0 to 1000 rpm, $HAuCl_4$ concentrations of 6.8, 12.5, 25, 50, and 100 mM, and deposition duration of 100 and 300 seconds.

First, the electrolyte stirring level during electrodeposition was investigated. To reduce the impact of diffusion on the rate and heterogeneity of electrodeposition for a given electrode size, increasing agitation allows to enhance replenishment. We observed that without agitation, uniform nanostructuring of the surface was impossible, even for the most concentrated solution such as 50 and 100 mM $HAuCl_4$. By inducing stirring, the thickness of the diffusion layer formed at the proximity of the electrode decreases, and the ion reduction rate increases. Different rotation speeds were tested, and it was determined that 1000 rpm was the best achievable rotation speed for electrolyte renewal close to the electrodeposited surfaces (see Figure S3, *Supplementary Data*). For slower rotation speeds, the average deposition current was lower, as the deposition rate was limited by the insufficient diffusion of ions near the electrode. These observations were correlated with inhomogeneous structuring.

<figure><img src="image_5.png" /><figcaption>Figure 2 - Impact of deposition parameters on nanostructure morphologies and dimensions. Scale bar is 1 µm. A. SEM micrographs displaying the nanostructures obtained at the center of the electrodeposited surface for 25, 50 and 100 mM $HAuCl_4$ concentrations and two deposition durations (100 s and 300 s). Spikes and scales are observed, and the average height of the nanostructures measured on AFM images is displayed in the top left corner of every micrograph (averaged on one batch). B. Photographs and SEM micrographs displaying nanostructure shape variations between the center and the edge of an electrodeposited sample for two gold-salt concentrations. The sample on the left column was nanostructured using an electrolyte of 6.8 mM of $HAuCl_4$ with 1 mM $Pb(CH_3COO)_2$, with a deposition duration of 540 s. The sample on the right column was made using a 100 mM of $HAuCl_4$ with 1 mM $Pb(CH_3COO)_2$ during 100 s.</figcaption></figure>
Second, the deposition duration was investigated. Two reference durations of electrodeposition were tested (100 seconds and 300 seconds) for different $HAuCl_4$ salt concentrations, in batches of five microscope slides (see Figure 2). The SEM and AFM results highlighted that by increasing deposition duration, nanostructures obtained for the same concentration had larger heights and widths. For example, the nanostructures created using 100 mM $HAuCl_4$ had a mean height increase from 220 ± 16 nm to 493 ± 16 nm. By increasing the deposition duration, it allowed for the accumulation of electrodeposited matter and thus increased nanostructure height and width, while decreasing spike density. As the surface undergoes different nucleation steps, several morphologies, such as nanospikes and nanoscales, could also be observed (see Figure 2B). Spikes are conical nanostructures with an aspect ratio over two, ending with a tip of only a few nanometers in diameter and a larger base (100 - 300 nm). Overall, spatially uniform deposition of defined spikes with heights ranging from approximately 71 to 575 nm was obtained for $HAuCl_4$ concentrations larger than 25 mM and deposition durations longer than 100 seconds.

Finally, the impact of $HAuCl_4$ concentrations in water on the electrodeposited nanostructure geometry was investigated. The 6.8 and 12.5 mM concentrations did not allow for obtaining uniform nanostructure on the 12 cm² substrate and large variations in average spike height and density between the edges and the center of the sample was observed. For 100 seconds electrodeposition duration, the overall roughness of the surface increased from 0.6 nm (considered as smooth gold) to 8 nm for 6.8 mM $HAuCl_4$ concentration and 28 nm for 12.5 mM $HAuCl_4$ concentration. However, no well-defined nanostructures could be observed using SEM and AFM, especially in the center of the sample. By increasing deposition duration to 540 seconds, some nanostructures were observed on the edge of the nanostructured sample, with spikes of aspect-ratio over three. The structuration only extended a few microns towards the center of the sample (see Figure 2A, left). For the rest of electrodeposited area on the microscope slide, the size of gold atom clusters increased, increasing the roughness, but the surface remained without define nanostructuration. By progressively increasing $HAuCl_4$ concentration to 25, 50, and 100 mM, the uniformity of the deposition over microscope slides was improved. SEM images taken at different locations of the sample showed identical structuration in the center and near the border of the sample surface (see Figure 2A, right). These findings were confirmed by AFM measurements, with an average coefficient of height variation under 10% per slide for 100 mM $HAuCl_4$ electrodeposition. Regarding the morphologies of the nanostructures obtained, different geometries were observed for the three concentrations, both nanospikes and nanoscales (see Figure 2A, 2B). These nanostructures were mostly observed for concentrations such as 25 mM and 50 mM and 100 seconds deposition durations. For higher concentrations and deposition durations, nanostructures mimicking sharp fish scales were visible. Scales are tetragonal pyramidal nanostructures, with greater growth along only one axis and are often mixed with some spike-like nanostructures (see Figure 2A, 100mM).

Overall, a minimal $HAuCl_4$ concentration of 25 mM was established as the threshold for the uniform electrochemical nanostructuring of 12 cm² large samples. This $HAuCl_4$ concentration threshold was confirmed even by increasing deposition duration. For 800 seconds deposition duration, homogeneous nanostructuration could not be obtained for 3.4, 6.8, and 13.6 mM $HAuCl_4$ concentrations (data not shown). Interestingly, in the literature, such low $HAuCl_4$ concentrations have previously been used to successfully create gold nanostructures.[10,11,14]
However, the electrodeposited surface areas used in these studies were microelectrodes, with much smaller surfaces to be nanopatterned (16 mm² discs for Sabri *et al.*, for example).[10]

The nanostructuration differences obtained for the lowest concentrations compared to the literature can be explained by the size of the electrode. Micro-electrodes operate in a regime where diffusion fields are small and ion transport is efficient and thus not limiting. At lower concentrations, mass transport is less likely to be a limiting factor compared to macro-electrodes. The thickness and homogeneity of a thin film are directly dependent on the deposition rate, which is proportional to the current density, i.e., the number of electrons passing through the electrode. According to Faraday's law, the total current *I* is proportional to the electrode surface area *A* and the molar flux of the electroactive species. Under diffusion-limited conditions, this flux can be described by Fick's law, which relates it to the diffusion coefficient *D*, the bulk concentration *C*, and the diffusion layer thickness *δ*. By combining these two laws, the expression for the diffusion-limited current becomes:

$$I_{\text{lim}} = nF\frac{\text{DC}}{\delta}A \quad (1)$$

where *n* is the number of transferred electrons and *F* is the Faraday constant. This equation emphasizes the linear dependence of the limiting current on both the electrode surface area and the electrolyte concentration, highlighting the critical role of mass transport in electrochemical processes.

During electrodeposition, ion depletion occurs close to the electrode surface. This depletion is counterbalanced by ion transport by diffusion, migration and convection from the bulk solution. At low concentrations, the thickness of the diffusion layer *δ* is significant, leading to a decrease in *I*<sub>lim</sub>. Under these conditions, local heterogeneity can be observed. At the edge of the electrode, where the solution is essentially renewed, the deposition rate is higher, while in the middle of the electrode, local depletion zones induce a lower deposition rate. This effect is reinforced by the size of the electrode. Consequently, larger electrode surfaces require more concentrated electrolyte solutions to ensure adequate ion supply and enable the formation of uniform nanostructures. *I*<sub>lim</sub> increases proportionally to the concentration *C*, ensuring a steady ion flux to the entire electrode. The diffusion layer thickness *δ* becomes also thinner, reducing mass transport limitations. Additionally, the solution at the surface edges is always more efficiently renewed than at its center. At low electrolyte concentrations, this leads to heterogeneous deposits, with nanostructures forming predominantly at the edges of the sample.
<figure><img src="image_6.png" /></figure>

<figure><img src="image_7.png" /></figure>

<figure><img src="image_8.png" /></figure>

Figure 3 – Demonstrating the relationship between surface charge density and average peak height using AFM and deposition current measurements. A. Real time monitoring of the effect of HAuCl₄ concentration on the deposition current at a 0.05 V potential/Ag/AgCl. B. AFM imaging used to determine the rms and average height of the nanostructures created after 100 s electrodeposition of 100 mM HAuCl₄ at 0.05 V. C. Correlation between surface charge density after the electrodeposition and the average height of the nanostructures formed. The linear trend line is calculated based on more than 100 independent experiments. Each experimental point is illustrated with a symbol for the HAuCl₄ concentration (x - 100 mM; ■ - 50 mM; ♦ - 25 mM; ● - 12.5 mM and ▲ - 6.8 mM) and a color for the deposition duration (Dark green - 800 s; navy - 300 s; purple - 200 s; orange - 100 s and yellow - 50 s). For each group of identical parameters, the average height and surface charge density value are calculated and plotted with their 95% confidence interval (o).

As shown in Figure 3A, the current monitoring during electrodeposition validates Faraday’s first law, where increasing the electrolyte concentration results in a higher plateau value for the deposition current. Although equation (1) illustrates the relationship between current, electrode surface area, and electrolyte concentration, it does not fully explain the trends and values observed in the current curves. Specifically, the increase in current and deposition speed over time at higher concentrations, particularly 25, 50 and 100 mM, likely reflect secondary effects. Since these electrodepositions are performed under stirring conditions and over extended periods, the process is likely not diffusion limited. These secondary events may arise from the rapid
nucleation of structures at high concentrations or the geometry of the electrochemical cell. Current density measurements further confirm that increasing the electrolyte concentration to 100 mM HAuCl₄ results in a higher deposition rate, facilitating greater structuring over the same duration. SEM and AFM imaging corroborate these findings, showing larger and taller nanostructures at higher concentrations for the same electrodeposition time.

### 3.1.2. Nanostructure growth monitoring and morphological characterization

During this series of experiments, nanostructures were mainly characterized by average height of the nanospikes using AFM measurements (see Figure 3B). By correlating this parameter to the total surface charge density after deposition, a linear relationship between both measurements was established (see Figure 3C). The surface charge density was extracted from the electro-active area from the CV post-electrodeposition (see Figure S4, Supplementary Data) and the current density monitored during the electrodeposition process (see Figure 3A) using the following equation:

$$ \sigma = \int_{0}^{T} I dt \quad (2) $$

where I is the deposition current density in mA/cm² and T is the deposition duration in seconds. As previously highlighted, the deposition current directly depends on the HAuCl₄ concentration, and σ is obtained by integrating I over the deposition duration. Equation (2) clearly highlights that the surface charge density is dependent on these two deposition parameters. By linking both the HAuCl₄ concentration and the deposition duration with the surface charge density through the linear correlation, it is thus possible to predict the height of the electrodeposited nanospikes with great confidence (R = 0.96). Conversely, these results enable rapid estimation of key electrodeposition parameters for synthesizing nanostructures of a desired height and control of deposition using current monitoring (see Figure S5, Supplementary Data). However, the surface charge density is specific to each deposition setup, sample surface, as well as the electrolyte and electrodes used. Metal growth is dictated by several mechanisms with often non-linear dynamics, along with the set-up size and geometry and can be greatly impacted by the process conditions.[28,36] The different nanostructure shapes – from spikes to scales – showed that growth did not occur only toward the vertical axis, but also along other spatial directions. Then, for both the monitoring of gold salt concentration and stirring speed effects, recording of surface charge density on electrodeposited nanostructures added valuable information to the nanofabrication process.

### 3.1.3. Fabrication and characterization of 150 nm high gold nanospikes

After investigating the effects of deposition parameters on the average height of nanostructures, optimal parameters were selected to produce a series of 130 - 190 nm high nanostructured samples. These nanostructures are particularly relevant thanks to their dimensions similarity to nanospikes found on antimicrobial dragonfly wings.[37–39] The correlation established between nanostructure height and deposition conditions indicated that nanostructures of approximately 150 nm in height can be obtained with three different sets of experimental conditions: 100 mM HAuCl₄ concentration for 100 seconds, 50 mM HAuCl₄ concentration for 115 seconds, or 25 mM HAuCl₄ concentration for 150 seconds (see Figure S5). Ultimately, 100 mM HAuCl₄ concentration electrodeposited for 100 seconds were selected as the optimal
parameters to fabricate samples with the expected dimensions, in the shortest time and with the best homogeneity.

Morphological parameters of the electrochemically nanostructured samples were characterized using SEM and AFM imaging (see Figure 4). During the optimization of electrodeposition parameters, multiple independent batches were produced using the same deposition conditions. The average height of the nanostructures was calculated from at least three AFM images taken for multiple samples across these batches. (see Table S6, Supplementary Data). This approach, combined with the exclusion of surface extremities from testing, achieved by applying a self-adhesive silicone chamber, gave us confidence in the homogeneity of the nanostructures on the surface. While the electrodeposition process inherently leads to spikes of varying heights, some variation is expected. However, our process successfully minimized this overall variability across the surface. While other morphological parameters, such as interspacing, aspect ratio, or pitch, may play a more critical role in optimizing nanostructure antibacterial efficiency, the current literature does not provide a clearly identified optimal parameter.[40] Rather than focusing solely on optimizing surface topology control, our aim was to demonstrate both the feasibility of producing large-scale surfaces applicable to real-world production needs and the relevance of the characterization methodology on electroplated surfaces, which naturally yield heterogeneous topographies. While ordered nanostructured patterns are useful for verifying mechanistic hypotheses, heterogeneous surfaces are arguably more representative of real materials and may even enhance antibacterial performance, as reported in several studies.[9,41] This highlights the value of applying standardized characterization approaches to surfaces with varying nanometric topology.

A

<table><thead><tr><th>Parameters</th><th>Height (nm)</th><th>RMS (nm)</th><th>Density (µm<sup>-1</sup>)</th><th>Ratio S<sub>a</sub></th></tr></thead><tbody><tr><td>[Au] = 100 mM,<br/>[Pb] = 0.5 mM,<br/>Time = 100s</td><td>220 ± 16</td><td>43.6 ± 3</td><td>21 ± 2</td><td>2.9 ± 0.1</td></tr></tbody></table>

B

<figure><img src="image_9.png" /><figcaption>Figure 4 - Quantification of the nanospikes dimensions on a batch of five slides. A. The values of average height of the nanospikes, RMS, spike density and active surface area (Sa) are averaged from three different nanostructured samples in the same deposition conditions. The surface area (Sa) ratio is measured using Cyclic Voltammetry (CV) before and after the electrochemical deposition. B. SEM imaging confirmed the uniformity of the nanostructures on the sample and the spike density is averaged on three SEM images of x 15 000 magnification. The average height and RMS are measured using AFM. Scale bar is 2 µm.</figcaption></figure>
To ensure that any detected antibacterial activity resulted directly from the nanostructures, it was verified using X-ray photoelectron spectroscopy (XPS) and energy dispersive spectroscopy (EDX) that no residual Pb²⁺ or Cl⁻ were present on the surfaces after electrodeposition and rinsing (see Figure S7 and S8, *Supplementary Data*). The results showed no presence of these elements, which confirms previous observations. Several studies have specifically investigated the presence of unwanted precursor ions, such as Pb²⁺ and Cl⁻, in electrodeposited gold nanostructures, even when using significantly higher concentrations of lead salts than in our formulation (up to 2 mM for low concentration of Au³⁺). In particular, Elbourne *et al*. confirmed through XPS, EDX, and XRD analyses that no trace of Pb²⁺ or Cl⁻ was detectable on the surface of gold nanospikes following electrodeposition, supporting the conclusion that such ions do not remain on the surface after fabrication and therefore do not interfere with the surface properties or antibacterial effects of the nanostructures. This can be explained by the electrodeposition parameters, which allows Pb to be replaced by Au due to the higher deposition potential of Au, leaving no Pb on the surface of the Au nanospikes. [10,11,14]

## 3.2. Antibacterial activity assessment of the electrodeposited nanostructures

### 3.2.1. Critical overview of existing methods used for antimicrobial surfaces activity characterization.

Although surface contamination by bacteria has been a recognized problem for years, the development of antimicrobial surfaces is a relatively recent advancement. [42] While robust methods for characterizing antimicrobial agents in solution exist, standardized protocols for antimicrobial surfaces are still in development. This gap complicates the comparison of results across different studies and slows down the development of a comprehensive understanding of antimicrobial surfaces mode of action.

The evaluation of antibacterial surface efficacy relies on some standardized testing methods to ensure consistent and reliable results, used in both academia and industry. Among these, ISO 22196 and JIS Z 2801 reference methods utilize CFU counting as their main characterization method, where bacteria are applied to a surface, incubated, and then counted to determine the number of viable cells able to form colonies after plating. [43,44] ASTM E2149 reference method, on the other hand, uses a dynamic approach where the test material is shaken in a bacterial suspension, with CFU counts taken to assess bacterial reduction over time. [45] ASTM E2180 reference method also employs CFU counting but is specifically designed for materials with embedded antimicrobial agents. [46] In addition to CFU counting, fluorescence-based assays are often used alongside these methods, employing dyes to distinguish between live and dead bacteria, offering a more nuanced understanding of bacterial viability. [47] While these methods are widely used, they have limitations in replicating real-world conditions and often ignore the multi-effect action that surfaces can display, which can lead to variation in the efficiency measured using different methods. [48]

CFU-based characterization methods are relevant for studying antimicrobial surfaces using bactericidal agent release; however, the conclusions that can be drawn are limited for other modes of action. For surfaces without bactericidal effect, a decrease of bacterial concentration in the liquid media can be caused by the strong adhesion of the cell (dead or alive) on the surface. In the same manner, anti-fouling surfaces may not display a reduction in the viability of the cells in suspension, although this non-killing effect might be interesting for some antimicrobial
applications. To evaluate the different effects of an antimicrobial surface, it is thus necessary to quantify both the number of bacteria on the surface and in the medium after incubation.[47,49]

Fluorescent labeling can provide valuable information regarding the adhesion and viability of the bacteria adhered to the antimicrobial surface. Notably, the Live/Dead Baclight Kit™ is widespread in industrial and research protocols. This kit uses a combination of two fluorescent dyes that bind to nucleic acids, SYTO® 9 and propidium iodide (PI). SYTO® 9 permeates through the membrane of bacteria and fluoresces in green, whereas PI only enters and labels in red cells that have a damaged membrane, by displacing SYTO® 9 fluorophore due to its greater affinity toward nucleic acids. However, membrane integrity markers are not reliable viability markers, as they can label dead bacteria, as well as bacteria with reversibly damaged membranes. Thus, stressed bacteria with partially damaged membranes, but viable once detached from the surface (bacteriostatic effect), are often falsely labeled as dead ones (bactericidal effect). The overestimation of the dead bacteria ratio is often associated with PI staining and does not represent the real viability of bacteria in contact with the surface.[50,51] Another limitation originating from the Live/Dead kit is the double staining of bacteria. SYTO® 9 and PI staining of nucleic acids is a dynamic process, with the replacement of the SYTO® 9 dye by PI when the cell membrane is damaged. Thus, it is possible to observe some yellow or orange staining when both dyes are present in a bacterium. Most studies consider such cells as dead ones, thus overestimating the total dead bacteria count. Due to the limitations of this technique, it is hard to draw precise conclusions based on a single-step labeling experiment regarding the “real” viability state of the bacteria after being in contact with nanostructured surface without further examination.[52–54] It was thus concluded from previous studies that viability labeling results should always be confirmed by another method such as solid or liquid enumeration.[51] Nevertheless, to this day, fluorescence single-step labeling experiment remains one of the most trusted and widespread methods to evaluate adhered bacteria viability on surfaces. However, its cost and technical requirements may limit accessibility, particularly for smaller industrial laboratories that rely on standardized, lower-cost assays for regulatory approval.

To evaluate the viability of the cells after being in contact with a nanostructured surface, multiple methods consist in detaching bacteria from the surface to release them into the bulk liquid medium, followed by a solid medium enumeration of the bacteria detached in the liquid. In the literature, ultrasound baths have been extensively used for that purpose.[47,55,56]. However, it has been shown that they often are not reliable and the ultrasounds themselves can affect bacteria viability.[12,13] In fact, the effects of ultrasound on bacteria is highly impacted by various parameters, such as the frequency and power of the ultrasound apparatus used, the sample position in the bath, the duration of the ultrasound, etc. It has also been shown that the use of sonication impacts bacteria viability in a positive or negative manner depending on the ultrasound frequency and power, thus distorting the results of the solid enumeration.[12,13,57] Other methods consist in transferring bacteria by contact onto Petrifilm® or solid culture to enumerate the organisms. However, these methods are limited to the use of low concentrations (150 CFU/mL) and volume of bacterial suspensions (< 1 mL) and can be difficult to reproduce and use on different surfaces.[58]

To characterize nanostructure-induced antibacterial properties, a protocol inspired by Parreira *et al.*[59] based on a combination of adhesion and proliferation tests is designed, with optimized steps for the sample conditioning, bacteria culture, parallel characterization, solid enumeration, and fluorescent labeling (see Fig.
ure 5). For each assay, two identical nanostructured microscope slides are prepared and processed in parallel. One slide is used to evaluate the adhesive and contact-killing properties of the nanostructures, while the other slide is used to assess the ability of adhered bacteria to proliferate in a planktonic phase and form colonies on the nanostructured surface.

<figure><img src="image_10.png" /><figcaption>Figure 5 – Step by step illustration of the characterization protocol. The viabilities of planktonic, adhered on surface and released bacteria are assessed using two slides and a combination of fluorescence labeling and spot plating assay.</figcaption></figure>

First, to verify that the antibacterial effects observed are only due to nanostructured surfaces interaction with bacteria, an extensive cleaning protocol was used before and after electrodeposition, and sterilization of the surfaces was performed before the microbial characterization (see Figure 5, step 1). Then, the bacterial inoculum was prepared (see Figure 5, step 2), with bacteria in their exponential growth phase. In the literature, it has been shown that *Staphylococcus aureus* and *Pseudomonas aeruginosa* are differently impacted by nanostructured surfaces depending on their stage of growth.[9] In addition, sampling bacteria in their exponential growth phase allows studying the efficiency of antibacterial nanostructured surfaces in the most favorable conditions for bacterial development and with a highly active bacterial population. After the 2h incubation of the bacteria PBS (see Figure 5, step 3), the viability of planktonic cells exposed to the nanostructured surface is evaluated by using a spot plating assay (see Figure 5, step 4). The ability of bacteria to bind to nanostructured samples was assessed by counting the number of adhered bacteria using optical fluorescence microscopy and the viability state of these adhered bacteria was characterized using Live/Dead fluorescent labels (see Figure 5, step 5). The second slide is washed with PBS to retain only tightly bound bacteria (see Figure 5, step 6). Subsequently, tryptic soy broth (TSB) is used as a rich culture medium, favoring the bacterial growth and is added to the substrate (see Figure 5, step 7). After 4 hours of incubation at 37°C, the concentration of bacteria released from the nanostructures into the planktonic phase is measured, along with the counting and viability assessment of bacteria adhering on the nanostructured slides (see Figure 5, step 8 and 9). Incubation periods of 2 hours in PBS and 4 hours in TSB were selected to balance bacterial colonization of the surface for effective
fluorescence detection, while minimizing biofilm formation. This approach aims to facilitate the characterization of early-stage biofilm adhesion and growth inhibition mechanisms and thus, focus on short incubation durations only.

There are multiple advantages to this protocol compared to those used in the literature so far. The first advantage lies in the combination of multiple well-known techniques to reinforce the results obtained. By combining fluorescent labeling and solid CFU enumeration, it is now possible to quantify both the number of bacteria adhered to the surface and the number of bacteria still in planktonic form. This characterization allows elucidating the different mechanisms of action that an antibacterial surface can display, such as bactericidal by release, antibacterial by contact, or anti-adhesive. However, to discriminate a contact-killing surface from a bacteriostatic one, it is necessary to confirm the viability state of the bacteria in contact with the surface. This requirement leads to the second advantage of this protocol. The use of a second incubation step in rich medium with live/dead fluorescent labelling and CFU characterization allows the study of adhered bacteria without affecting their viability, unlike in ultrasound-based methods. This step allows to conclude whether the results observed in fluorescence are either due to a bacteriostatic or a bactericidal effect: if the red-labeled bacteria can reproliferate in the medium after detachment, the antimicrobial activity of the nanostructures is more bacteriostatic than bactericidal. This proliferation test, combined with a second fluorescence and spot plating assay, also allows verifying if the adhered bacteria can stay adhered (self-cleaning or not) and if they keep the ability to divide and grow on the surface and potentially form biofilm. Another critical aspect of the characterization of antimicrobial surfaces is the biological variability that can occur between repetitions of the protocol. To ensure reliable replication, and that characterization assays are carried out in similar standard conditions, silicon culture chambers are added onto nanostructured slides: multiplexed incubation wells enabled compartmentalizing the nanostructured surface and thus parallelizing the characterization, up to six times, on the same nano-sized motifs. Finally, the use of a spot plating assay allows covering a large range of dilution, as the viability of the bacteria in solution can drastically change when in contact with different antibacterial surfaces. It also enables the enumeration of the number of bacteria viable in solution more efficiently than standard counting by reducing the number of solid media to prepare and inoculate.[16,60–62]

Overall, using the adhesion and proliferation tests, combined with live/dead staining and spot plating assay, allow us to characterize multiple antibacterial properties from one single experiment, without interfering with the bacteria viability. This method could be adapted to assess promising emerging antibacterial and -microbial strategies with different modes of action, such as photoactivated surfaces using titanium dioxide, thereby helping to generalize findings on cell/surface interactions.

### 3.2.2. Effects of gold nanospikes on *Escherichia coli* adhesion, viability and reproliferation

The antibacterial properties of the 150 nm high nanospikes are assessed in comparison to non-nanostructured gold surfaces (CVD deposited gold on microscope slides) with two bacterial strains. These two strains were chosen because of their implication in biomaterials-related infections and their morphological and physiological differences.[63,64] Since motility and bacterial membrane thickness are two independent parameters known to impact antibacterial efficiency, we chose to test the surface efficiency against a motile Gram negative bacterium (*Escherichia coli*) and a non-motile Gram positive bacterium (*Staphylococcus epidermidis*).[30]
The results of antibacterial tests for *Escherichia coli* are illustrated in Figure 6, along with characterization of the live planktonic bacteria (Figure 6A), as well as counts of cells bound to surfaces and live/dead ratio (Figure 6B) after 2h incubation in PBS. After reincubation in rich medium, the numbers of live planktonic bacteria (Figure 6C) and surface-bound bacteria (Figure 6D) are also investigated. The detailed enumerations for *Escherichia coli* are available in Figure S9 of *Supplementary Data*.

<figure><img src="image_11.png" /><figcaption>Figure 6 – Antimicrobial effects of gold nanospikes on Escherichia coli for unbound bacteria in the 300 µL solution (A, C) and for bound bacteria on the surface (B, D). A. Spot plating assay after 2h in PBS. The blue line corresponds to the inoculum concentration and the band to its confidence interval. B. Fluorescence labeling quantification after 2h in PBS. C. Spot plating assay after 4h in TSB. D. Fluorescence labeling quantification after 4h in TSB. †The error bars for A and C represent the confidence interval (CI) at 95% on three experiments. For B and D, the cells labeled by PI only are in red and considered “Dead”; The cells labeled by SYTO® 9 only are in green and considered “Alive” and the cells labeled simultaneously by SYTO® 9 and PI are in orange and considered “Damaged”. Each point represents one quantification and is given to illustrate the natural variation in bacteria repartition on the surface. Statistically significant differences are indicated as *p* < 0.05, **p** < 0.01, ***p** < 0.001, ****p** < 0.0001 and NS, not significant (non-parametric Mann-Whitney test).</figcaption></figure>
After the first step of adhesion in bacteriostatic medium (Figure 6A), the enumerations of *Escherichia coli* planktonic cells show a significant difference in bacteria concentration of 36% between bare gold and nanospikes (p = 0.0002). A significant reduction of 13% compared to inoculum concentration is noticeable for live planktonic bacteria in contact with non-structured gold (p = 0.031) and a non-significant elevation of planktonic cells level compared to the reference is observed for bacteria in contact with nanostructures (p = 0.056). The change in bacterial concentration on surface between bare gold and nanostructures could be the result of a lower adhesion of the bacterial cells on the structured surface, which needs to be confirmed by surface imaging.

Fluorescence is then used to characterize the live state of the bacteria cells bound to the surface to confirm if the increase of viable cells in solution for the nanostructured gold surface is due to bacteria weak adhesion to the surface. In Figure 6B, bacterial adhesion values correspond to the sum of alive, dead, and damaged bacteria counts. The fluorescence image analysis shows that the total bacterial adhesion count on the surfaces after 2 hours incubation seems to be similar for both smooth and nanostructured surfaces. A minor decrease in the viable bacteria ratio (labeled in green) can be observed for the nanostructured surfaces, with more bacteria marked with both fluorescent labels (p = 0.10). This could be due to changes in *Escherichia coli* membrane integrity caused by the nanospikes. These results do not align with the low adhesion hypothesis, however the greater proportion of live bacteria adhered to the reference compared to the nanostructures could explain the decrease in live planktonic bacteria.

After the characterization of antibacterial effects on planktonic cells and cell adhesion, bacteria still bound to nanostructures after light washing are incubated in a rich culture medium to assess their ability to proliferate on the surface and/or to be released in a planktonic phase. Following the 4-hours incubation, the enumeration of live cells in the liquid medium (Figure 6C) showed +125% higher release of *Escherichia coli* from the nanostructured surfaces, with an average of 7.7 ± 0.3 × 10⁵ bacteria/well, compared to the non-structured gold surface (p = 0.00002). As the number of bacteria bound was similar for both surfaces after the initial incubation step, we can conclude that a larger number of bacteria are able to detach from the nanostructures compared to the reference and proliferate in the liquid medium. Fluorescence labeling after 4-hours incubation is then performed on attached bacteria (Figure 6D and Figure S10-A, Supplementary Data). For the non-structured gold substrate, an average of 2.5 × 10⁴ bacteria/well are enumerated, with 59% labeled viable, 38% in a damaged state, and only 3% labeled dead. On the nanostructured surface, an average of 1 × 10⁴ bacteria/well are enumerated, with 45% labeled viable, 43% in a damaged state and only 12% labeled dead. These results indicate a significant reduction of 61% in the total adhered bacteria on the nanostructured surface compared to non-structured gold substrate (p = 0.043). This difference is mainly caused by the significant decrease of 65% of viable bacteria on the nanostructured surface (p = 0.005). Surprisingly, the combination of both characterization techniques indicates that *Escherichia coli* cells tend to proliferate more easily in planktonic form when released from gold nanostructures than after incubation on the control non-structured surface. Since bacterial adhesion is similar to non-structured gold substrate and nanostructured gold substrate after the initial 2-hour adhesion step, but a larger number of bacteria are counted in solution after reincubation for nanostructures, along with a decrease in total bacterial adhesion compared to smooth gold, it can be concluded that bacteria detach more easily from the nanostructured surface. The same characterization protocol is applied to 400 nm nanostructures
and displayed similar results (see Figure S11, Supplementary Data). This indicates that the nanostructures' efficiency is more dependent on other nanostructure geometric features (tip radius, aspect ratio, nanostructures density, etc.) rather than on nanostructures' average height on the range tested.

Overall, these results highlighted a reduction in *Escherichia coli* growth on nanopatterned surface compared to unstructured surfaces used as controls. These results are significantly different from previous studies found in the literature, but it can be explained by the methodological differences and the nanostructures' shape variations.[11,65] In the study by Wu *et al.*, the nanostructures' shapes are rings, pillars, and nuggets, and the reproliferation tests are based on OD measurements and not fluorescence and CFU counting, while Elbourne *et al.* used only fluorescence with longer incubation time (18h) and no reproliferation tests.

In the current study, it is important to highlight that the antibacterial effect of the nanostructures is tested in solution, with agitation, and that the surfaces are not dried at any point. The mechano-bactericidal effects of nanostructures are still under investigation and it has been postulated that the air/liquid interface could play a major role in nanostructure efficiency.[66] By having bacteria at the air/liquid interface, the antibacterial property of nanostructured samples is increased due to the surface tension, applying a normal force on the bacteria, increasing deformation and leading to membrane rupture.[67] In contrast, during drying or evaporating droplets, bacteria are exposed to additional physical forces that drive their accumulation at the solid-liquid-air interface, a phenomenon known as the coffee-ring effect. This can influence surface antibacterial efficacy, as bacterial cells may stack on top of each other upon drying and only the cellular monolayer directly in contact with the surface is inactivated, while cells lying above them would remain viable. Considering such interface-driven effects is therefore essential when comparing results across different experimental setups.[68]

### 3.2.3. Effects of nanospikes on *Staphylococcus epidermidis* adhesion, viability and reproliferation

Effects of the 150 nm high gold nanostructures are tested against *Staphylococcus epidermidis* bacteria (detailed enumerations are available in Figure S12 of Supplementary Data). After incubation on planar gold substrate and nanospikes in bacteriostatic medium, the enumeration results showed no significant difference in bacterial liquid suspension concentration (see Figure 7A). Total bacterial adhesion counts highlighted similar adhesion for both surfaces (see Figure 7B and Figure S10-B, Supplementary Data), and a similar viable cells proportion on the surfaces. The fluorescent labeling also showed that the mortality of bacteria (labeled in red) has more than doubled for nanostructures (152% increase, p = 0.04), with 48% labeled dead cells, compared to counts on non-structured gold substrate. Combined to the significant damaged cells population (in orange) decrease for nanospikes, the results could indicate a bacteriostatic or bactericidal effect by contact on the nanostructured substrate. Then, *Staphylococcus epidermidis* still attached after gentle rinsing on both surfaces are reincubated in a rich culture medium. Following the 4-hours incubation at 37°C in a rich culture medium (see Figure 7C), the planktonic cell enumeration estimates at 2.25 ± 0.5 × 10⁴ the number of bacteria per well for the nanostructured surfaces, which is significantly fewer than the 3 ± 0.1 × 10⁴ viable bacteria for the non-structured gold substrate (27% reduction, p = 0.01). This could be explained by a lower release of *Staphylococcus epidermidis* bacteria from the surface, or by the killing of bacteria adhering to nanostructured surfaces. However, the results of the fluorescent images counting (see
*Figure 7D*) showed fewer cells attached to the nanostructured surface than to smooth gold (69% reduction, p = 0.005). For bacteria attached to the non-structured gold substrate, an average of 1.2 × 10⁵ bacteria/well are enumerated, with 25% labeled viable, 73% in a damaged state, and only 1% labeled dead. For bacteria on the nanostructured surface, an average of 3.9 × 10⁴ bacteria/well are enumerated, with 82% labeled viable, 15% in a damaged state and only 3% labeled dead. The overall reduction of viable bacteria both in liquid phase and on the nanostructured surface showed that these gold nanostructures induced a significant contact-killing effect. The apparent ~50% killing observed by fluorescence imaging should be interpreted with caution, as a non neglectable proportion of bacteria did not adhered to the surface in the first place. The absence of fluorescent dead-labeled bacteria on the nanostructured surface after rinsing and 4 hours of incubation also reveals a self-cleaning effect, but could equally be explained by complete lysis of the bacteria after step 1, which would not appear in fluorescence monitoring.

<figure><img src="image_12.png" /></figure>

<figure><img src="image_13.png" /></figure>

<figure><img src="image_14.png" /></figure>

<figure><img src="image_15.png" /><figcaption>Figure 7 - Antimicrobial effects of gold nanospikes on Staphylococcus epidermidis for unbound bacteria in the 300 µL solution (A, C) and for bound bacteria on the surface (B, D). A. Spot plating assay after 2h in</figcaption></figure>
PBS. The blue line corresponds to the inoculum concentration and the band to its confidence interval.
B. Fluorescence labeling quantification after 2h in PBS C. Spot plating assay after 4h in TSB. D. Fluorescence labeling quantification after 4h in TSB. †The error bars for A and C represent the confidence interval (Cl) at 95% on three experiments. For B and D, the cells labeled by PI only are in red and considered "Dead"; The cells labeled by SYTO® 9 only are in green and considered "Alive" and the cells labeled simultaneously by SYTO® 9 and PI are in orange and considered "Damaged". Each point represents one quantification and is given to illustrate the natural variation in bacteria repartition on the surface. Statistically significant differences are indicated as * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001 and NS, not significant (non-parametric Mann-Whitney test).

The difference in antibacterial properties of the nanostructures on *Escherichia coli* and *Staphylococcus epidermidis* strains might be due to motility ability, which may play a greater role in reducing antibacterial effect on motile cells, rather than a protective effect due to the Gram-positive membrane thickness.[69] Indeed, *Staphylococcus epidermidis* cells, which are non-motile bacteria, seemed to be more sensitive to the 150 nm nanostructures than *Escherichia coli* bacteria, even though their thicker cell membrane could reduce the cell distortion and prevent mechanical rupture induced by contact with nanostructures.

## 4. Conclusions

Unraveling bacterial interactions with surfaces remains a major challenge due to the complexity of microbial behavior and biofilm formation. This article presents a thorough characterization of the antibacterial effects of surfaces, integrating adhesion and proliferation assays to provide a deeper insight into how nanostructured surfaces influence different bacterial activities on the nanostructured surfaces and detachment ability into the surrounding liquid environment. Using an ultrasound-free protocol, antimicrobial properties of gold electrodeposited nanostructured surfaces with 150 nm high features were characterized, highlighting differences in the effects on *Escherichia coli* and *Staphylococcus epidermidis* bacterial strains. After incubating surface-bound bacteria under proliferation-favoring conditions, a reduction in bacterial adhesion was observed for both strains, along with distinct differences in their proliferation behavior in liquid. Gold nanospikes clearly exhibited a contact-killing and self-cleaning effects with *Staphylococcus epidermidis*. Interestingly, results for *Escherichia coli* were less clear-cut, but still highlighted weaker interactions between bacterial cells and the nanostructured surface than with control planar gold surfaces. These variations in microbial behaviors could be attributed to microbial morphological differences, such as motility and cell wall structure. Further investigations with other biomedically-relevant bacteria, such as non-motile Gram-negative (ex *Shigella spp*) and motile Gram-positive (ex *Clostridium perfringens*) species, is warranted.

The study also explored various challenges in the development of antimicrobial nanostructured surfaces, focusing on the difficulty of producing uniform, large-area samples that meet application requirements. We demonstrated that electrodeposition is a viable method for producing bioinspired gold nanostructures on centimeter-sized surfaces, which is essential for statistically significant characterization. This method is cost-effective, allows rapid sample production, and provides tunable nanostructured surfaces by adjusting the electrochemical deposition parameters. Key values for uniform nanostructuring were identified by studying the effects of deposition time, HAuCl₄ concentration, and electrolyte stirring speeds. Surfaces up to 12 cm² were successfully structured at the nanometer scale with various nanosized morphologies, such as dragonfly-inspired nanospikes and fish-like nanoscales, with heights ranging from ~90 to
500 nm. A correlation between nanofeature height and deposition parameters was established, helping to predict nanostructure height or identify relevant deposition conditions, such as 100 mM HAuCl₄ for 100 seconds. Based on assays carried in model conditions, our study highlights the need for accurate, comprehensive characterization methods that take into account the potential for multiple effects of nanostructures on microbial developments on surfaces. Current literature on this topic often characterizes only one aspect of antimicrobial properties, such as adhesion after incubation or viability of planktonic cells, making it difficult to correlate results across studies. To advance the effective use of antimicrobial nanostructured surfaces, comprehensive characterization methods that account for multiple antimicrobial effects are essential.[64,65] Although this study focused on bacterial models, the proposed approach could be readily applied to other microorganisms, thereby contributing to a broader understanding of how nanostructured surfaces influence microbial colonization and survival. Consistent testing in conditions where samples may have been exposed to 'real life' conditions, meaning exposed to dust, soiling, heat or contaminant exposure, will also enable the creation of a robust knowledge base that will highlight global trends and elucidate mechanisms of action, such as the mechano-bactericidal effect.
## Acknowledgements

This work is a collaboration between the Laboratoire Nanotechnologies et Nanosystèmes (LN2), hosted by the Institut Interdisciplinaire d'Innovation Technologique (3IT) of Sherbrooke, and the Systèmes Moléculaires et nanoMatériaux pour l'Énergie et la Santé (SyMMES) hosted by the CEA Grenoble. LN2 is an International Research Laboratory (IRL) funded and co-operated by Université de Sherbrooke (UdeS), Centre National de la Recherche Scientifique (CNRS), École Centrale Lyon (ECL), Institut National des Sciences Appliquées de Lyon (INSA Lyon), and Université Grenoble Alpes (UGA). We would like to thank CMC Microsystems. We gratefully acknowledge the financial funding of the Biosurf Project by the CNRS 80|Prime, by Labex ARCANE, and by CBH-EUR-GS (Grant ANR-17-EURE-0003). We also acknowledge support from the National Science and Engineering Research Council (NSERC) of Canada through its Discovery Grants program (MC and PC). The authors also wish to thank G. Beaudin, R. Mathey, M. W. Hassen, Pr. P. Dauphin-Ducharme, and N. Fontaine for their scientific and experimental inputs.

## Declaration of generative AI and AI-assisted technologies in the writing process

During the preparation of this work the authors used Microsoft Copilot and DeepL Write tools in order to improve language and sentence structure. After using this tool/service, the authors reviewed and edited the content as needed and take full responsibility for the content of the publication.
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