Tailoring nanotopography and antibacterial properties of calcium phosphate bone grafts via fluoride incorporation

Carla Arca-Garcia; Maria Godoy-Gallardo; Maria-Pau Ginebra

doi:10.1016/j.bioactmat.2025.12.026

. 2025 Dec 27;59:205–223. doi: 10.1016/j.bioactmat.2025.12.026

Tailoring nanotopography and antibacterial properties of calcium phosphate bone grafts via fluoride incorporation

Carla Arca-Garcia ^a,^b,^c, Maria Godoy-Gallardo ^a,^b,^⁎, Maria-Pau Ginebra ^a,^b,^c,^d,^⁎⁎

PMCID: PMC12803813 PMID: 41541278

Abstract

Despite advances in bone graft design and surgical techniques, bacterial infection remains a major cause of graft failure, exacerbated by the global rise in antimicrobial resistance. This has intensified the pursuit of antibiotic-free strategies to prevent bacterial colonization. Among these, antibacterial surface nanotopographies have emerged as promising tools, leveraging nanoscale geometries to physically disrupt bacteria upon contact. In this study, we engineered the surface of a calcium phosphate bone graft to confer antimicrobial functionality through a dual approach: the creation of high-aspect-ratio nanotopographies and ionic doping with fluoride. Through controlled hydrolysis of α-tricalcium phosphate by biomimetic and hydrothermal treatments, we generated calcium deficient hydroxyapatite nanoneedle structures whose morphology and biofunctionality were tuned via fluoride incorporation. XRD and Raman spectroscopy confirmed the formation of hydroxy-fluorapatite, with phase composition and surface morphology dependent on fluoride concentration and processing parameters. Fluoride doping significantly altered nanoneedle dimensions and spacing and enhanced bactericidal activity, particularly against P. aeruginosa, and to a lesser extent S. aureus. Notably, fluoride-doping alone showed no antibacterial effects; however, when combined with nanotopography, a synergistic increase in efficacy was observed. Importantly, the antimicrobial surfaces supported the proliferation and osteogenic differentiation of SaOS-2 cells. Co-culture assays modeling pre- and post-implantation infection scenarios demonstrated robust cell adhesion and markedly reduced bacterial colonization. In conclusion, our findings present a multifunctional, synthetic bone graft with both physical and chemical antibacterial properties, offering a promising strategy to mitigate infection risks while supporting osteointegration.

Keywords: Antibacterial nanotopography, Calcium phosphate, Hydroxyapatite, Fluoride, Bone regeneration

Graphical abstract

Highlights

•
Fluoride doping enables tuning of calcium-deficient hydroxyapatite nanoneedle structures.
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Fluoride incorporation combined with nanotopography elicits a synergistic increase in antibacterial efficacy.
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Fluoride-doped nanoneedle surfaces significantly inhibit P. aeruginosa and reduce S. aureus metabolic activity.
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Co-culture pre- and post-implantation infection models demonstrate robust cell adhesion and reduced bacterial colonization.

1. Introduction

Calcium phosphates (CaP) are a family of inorganic compounds abundantly found in human bones and teeth. The principal mineral phase of bone is carbonated apatite [1]. This inherent affinity between CaPs and the human hard tissues underpins their widespread application as biomaterials for bone regeneration, particularly as synthetic bone grafts. Their exceptional biocompatibility, osteoconductivity and osteogenic properties make them ideal candidates as scaffolds for bone tissue engineering [2]. However, despite significant advances in surgical techniques, graft design, and material developments aimed at optimizing osseointegration and minimizing trauma or implant-tissue mismatch, complications related to bone grafts remain a significant clinical concern. These complications (e.g., delayed healing, graft failure and in severe cases, patient mortality), can arise from various factors, being bacterial infection recognized as one of the predominant causes of bone graft failure [3].

Infections related to bone grafts remain a challenging clinical issue [3]. They may arise either from intraoperative contamination during the surgical procedure or via hematogenous spread, wherein bacteria originating from distant sites within the body disseminate through the bloodstream and colonize the graft. The first few hours following implantation involve the so-called “race for the surface”, representing the competition of the graft surface between tissue cell integration and bacterial adhesion [4]. Preventing early bacterial attachment is critical for promoting successful tissue integration. However, this remains inherently challenging, as microorganisms possess a remarkable capacity to adhere to biomaterial surfaces and to establish long-term biofilms. Once mature, such biofilms provide long-lasting protection against antibiotics and facilitate evasion of the host immune response, allowing the microbes to persist on biomaterial surfaces for prolonged periods [5,6]. Therefore, effective strategies must focus on preventing early bacterial adhesion and disrupting early biofilm formation to ensure successful implant integration.

This challenge is further aggravated by the global rise of antimicrobial resistance (AMR), considered an emerging epidemic [7]. Of particular concern is the increasing prevalence of methicillin-resistant Staphylococcus aureus (MRSA), one of the most common causative agents of hospital-acquired infections [8]. Consequently, in the light of the escalating threat of antibiotic resistance, the development of antibiotic-free strategies to prevent bacterial colonization of biomaterials has emerged as a critical priority.

To address this challenge, various antibacterial surfaces have been developed through chemical modification with metals [9], antibacterial ions [10], phenols [11], and other antimicrobial agents [12]. These approaches typically depend on the sustained release of the active agents. However, their efficacy diminishes over time as the antimicrobial component becomes depleted. To circumvent this limitation, a promising alternative lies in contact-active antibacterial systems that eradicate bacteria through direct physical interaction. These systems employ biomaterials that incorporate high-aspect-ratio nanostructures capable of mechanically disrupting bacterial cell walls. Such nanoengineered surfaces create an environment that is inhospitable for bacterial adhesion, and any bacteria that do adhere are subjected to physical lysis. This lysis, is hypothesized to occur via two main mechanisms: either through mechanical stress from the stretching of the bacterial wall between adjacent nanoneedles, leading to rupture, or through direct perforation of the cell wall by the nanoneedles [13]. A key advantage of this approach is that the antibacterial effects are derived primarily from surface topography, irrespective of its chemical composition.

A variety of materials has been employed to fabricate antibacterial nanostructured surfaces, including silicon, titanium [14], and polymers [15]. For instance, Linklater et al. compared different polymer substrates with various nanotopography and showed that nanopillar arrays with a height of 60 nm and an inter-pillar spacing of 60 nm were able to disrupt the cellular integrity of both Staphylococcus aureus (S. aureus) and Pseudomonas aeruginosa (P. aeruginosa). However, several factors influence the overall bactericidal performance of these nanopatterns, including their physical characteristics, surface chemistry, and the specific bacterial strain involved [16]. Hence, these nanostructured surfaces offer a promising choice for providing long-term antibacterial activity on biomaterials. Our group has previously developed synthetic bone grafts based on low-temperature dissolution-precipitation reactions that mimic natural biomineralization [[17], [18], [19]]. This approach offers significant potential for fine-tuning the surface nanostructure of the resulting materials. In particular, calcium deficient hydroxyapatite (CDHA) substrates obtained by hydrolysis of α-tricalcium phosphate (α-TCP) can be tailored simply by modifying the reaction conditions, such as temperature, time, relative humidity or particle size [[20], [21], [22]]. Adopting this strategy, in a recent work we demonstrated the feasibility of engineering CaP materials with high-aspect-ratio nanotopographies exhibiting potent antimicrobial activity against Gram-negative bacteria, identifying surface morphology as the main factor contributing to bacterial death [23].

In this work, we aim to take a further step by leveraging the inherent substitutional flexibility of the hydroxyapatite crystal structure as a powerful tool for tailoring its properties and nanostructure [[24], [25], [26]]. The incorporation of doping agents offers a means to influence the morphology and aspect ratio of precipitated hydroxyapatite crystals. Among these, fluoride stands out as a particularly promising candidate, not only due to its role as a key component of biominerals such as dental enamel apatite [27,28], but also because it has been extensively studied for its ability to tailor the physicochemical properties of CaP materials, thereby enhancing their biomedical functions [28,29]. The present work explores the potential of ionic doping with fluoride with two main objectives. Firstly, we aim to use fluoride as a method to further modify and fine-tune the nanotopography itself, as previous studies have shown that fluoride incorporation in apatite can promote crystal growth along the c-axis [25,27,30]. Secondly, we explore whether the well-established intrinsic antibacterial properties of fluoride can act synergistically with the mechanical effects of the nanotopography, which would represent a relevant innovation for enhancing the overall antimicrobial efficacy. The fluoride ion is known to inhibit key metabolic enzymes such as enolase, involved in the glycolysis, thereby reducing ATP production and acid generation. Moreover, it interferes with proton export, leading to intracellular acidification and ultimately restricting bacterial growth and proliferation [[31], [32], [33], [34]].

Different nanotopographical features were obtained by the hydrolysis of α-TCP in fluoride solutions with varying concentrations under both autoclave (AC) and biomimetic (BM) environments. The two synthesis methods were chosen because it is known that by varying the conditions under which the hydrolysis of α-TCP to CDHA takes place, it is possible to control the size and morphology of the apatite crystals [35]. This provided the possibility of obtaining surfaces with the same composition but different nanostructural parameters, which may have an impact on their bactericidal properties. After thorough physicochemical characterization, antibacterial efficacies were assessed against two clinically relevant pathogens, namely S. aureus and P. aeruginosa. The cytocompatibility and inflammation response of the developed nanotopographies was evaluated with SaOS-2 preosteoblastic cells and RAW264.7 macrophages, respectively. Finally, the performance of the different nanostructured CaPs using either a pre-implantation or a post-implantation infection models was assessed. Overall, we demonstrated that engineered nanotopographies act synergistically with fluoride inherent antibacterial properties, resulting in an enhanced antimicrobial effect.

2. Materials and methods

2.1. Materials

Sodium fluoride (NaF), Triton X-100, Cytotoxicity Detection Kit LDH, calcium hydrogen phosphate (CaHPO₄) and calcium carbonate (CaCO₃) were purchased from Sigma-Aldrich (MO, USA). Commercially pure grade II titanium was purchased from Acnis International (France). McCoy's 5A Medium, Dulbecco's Modified Eagle Medium (DMEM), fetal bovine serum (FBS), L-glutamine, penicillin/streptomycin (p/s, 10 000 U mL⁻¹ and 10 000 μg mL⁻¹, respectively), TrypLE, Mammalian-Protein Extraction Reagent (MPER)®, Luria broth (LB), Brain heart broth (BHI), glycine (Gly), HEPES buffer solution, LIVE/DEAD BacLight bacterial viability kit, Thermo Scientific™ Pierce™ BCA Protein Assay Kits, hexamethyldisilazane (HMDS), DAPI and phalloidin-Alexa Fluor 546, UltraPure™ DNase/RNase-Free Distilled Water and SuperScript™ III Platinum™ SYBR™ Green One-Step qRT-PCR Kit were purchased from Fischer Scientific (MA, USA). Dulbecco's Phosphate Buffered Saline (PBS) was purchased from Capricorn Scientific (Germany). BacTiter-Glo Reagent was purchased from Promega (WI, USA). CellBrite™ Fix 555 was purchased from LabNet Biotecnica (Spain). Mouse inflammation antibody array (ab133999) was purchased from Abcam (UK). Paraformaldehyde (PFA) was purchased from ANAME (Spain). SensoLyte® pNPP Alkaline Phosphatase Assay Kit was purchased from Anaspec (CA, USA). RNeasy Minikit (250) was purchased from Qiagen (Germany). SaOS-2 cell line (HTB-85), RAW 264.7 cell line (TIB-71), Pseudomonas aeruginosa (ATCC-27853) and Staphylococcus aureus (ATCC-25923) were purchased from American Type Culture Collection (VA, USA).

The ICP solutions were made with ultrapure water (Milli-Q gradient A 10 system, resistance 18 MV cm, TOC <4 ppb, EMD Millipore, MA, USA).

2.2. Fabrication of nanostructured CDHA substrates

2.2.1. Powder preparation

To synthesize α-TCP, CaHPO₄ and CaCO₃ were combined in a 1:2 M ratio. The mixture was sintered at 1400 °C for 15 h, followed by air quenching at room temperature to stabilize the high-temperature α-phase. Subsequently, the α-TCP was milled as previously described to produce a powder with a median size of 2.8 μm [23]. In brief, the powder was milled for 40 min at 450 rpm using 10 agate balls (30 mm diameter, Serviquimia, Spain), followed by an additional 60 min at 500 rpm with the same agate balls. Finally, a further milling step was conducted for 60 min at 500 rpm using smaller agate balls (10 mm diameter).

2.2.2. Preparation of nanostructured CDHA discs

Disc-shaped specimens were prepared by uniaxial compaction of α-TCP powder using a manual hydraulic press (Atlas™ Manual 15T, Specac, UK). Two sizes were produced: 5 mm diameter discs were obtained by compressing 50 mg of powder at 1.5 tons for 2 min, while 10 mm diameter discs were prepared by applying 4.5 tons to 140 mg of powder for the same duration. In both cases, the resulting discs exhibited a uniform thickness of approximately 1 mm. All experiments were conducted using 5 mm diameter discs, except for phase characterization by X-ray diffraction (XRD), which was performed on 10 mm discs. To generate various nanoneedle topographies, the α-TCP discs were hydrolyzed to CDHA under different conditions (Fig. 1). Depending on the disc diameter, each sample was immersed in either 9.0 (5 mm) or 25.2 mL (10 mm) of solution to maintain a consistent ratio between mass of the sample and volume of the soaking liquid. The samples were submerged in either deionized water or NaF solutions of varying concentrations (0, 0.5, 1, 2 and 10 mM), and subjected to two distinct treatments: (i) incubation at 37 °C for 7 days (biomimetic treatment), or (ii) autoclaving at 121 °C and 2 atm for 20 min. Samples were coded as BM or AC to indicate biomimetic or autoclave treatment, respectively, followed by an F (indicating fluoride) and a number corresponding to the NaF concentration in the immersion medium (0, 05, 1, 2, or 10). Additionally, flat CDHA samples (coded as Flat) were prepared as controls by first hydrolyzing 5 g of α-TCP powder in 45 mL of deionized water at 37 °C for 7 days (biomimetic treatment), then drying and compressing the resulting powder using the previously described pressing protocol. Moreover, to independently evaluate the effect of fluoride without the influence of surface topography, control samples were fabricated by re-pressing the fluoride-doped materials to eliminate the nanostructure. Polished titanium discs (coded as Ti) were used as additional controls.

Fig. 1 — Schematic representation of the nanotopography synthesis process and evaluation procedures. i) α-TCP powder was first synthesized, and ii) either compressed into discs using a manual hydraulic press or iii) hydrolyzed to obtain CDHA powder. The α-TCP discs were subsequently subjected to autoclaving or biomimetic treatment, exposed to iv) fluoride-based solutions or v) water, to induce surface nanotopographies. vi) The CDHA powder was compressed to generate a flat surface of CDHA. Finally, vii) *in vitro* studies with bacteria and cells were conducted under mono- and co-culture conditions.

2.3. Physico-chemical characterization of the surfaces

2.3.1. Morphological analysis

The surface and cross-section of the discs were analyzed by scanning electron microscopy (SEM) with a Zeiss Neon40 FE-SEM (Carl Zeiss NTS GmbH, Germany) equipment. For each sample, five images were taken at a working distance of 4 mm and a potential of 5 kV. Previously, the discs were coated with carbon (10 nm thickness) using a high vacuum sputtering Leica EM ACE600 (Leica, Germany). For cross-sectional images, samples were coated using iridium (6 nm thickness). Image analysis was conducted to assess the distribution of needle length, spacing and density with the Fiji/ImageJ software (NIH, WI, USA) [36]. For height measurements 100 needles were counted, and for needle spacing and density 400 distances and needles were taken respectively.

2.3.2. Phase composition

Phase characterization was conducted on the 10 mm discs by XRD on a D8 Advance diffractometer (Bruker, MA, USA) equipped with a Cu Kα anode, operating at 40 kV and 40 mA. Data were collected in 0.02° increments over a 2θ range of 10°–60°, with a counting time of 2 s per step. Grazing incidence XRD was also performed on the same equipment and under the same conditions but with 3 s per step and configured at an incidence angle of 1°. The experimental diffraction patterns were compared with reference patterns for CDHA (PDF# 00-046-0905), α-TCP (PDF# 04-010-4348), and β-TCP (PDF# 04-008-8714). Phase quantification was obtained by Rietveld refinement using Profex software (Profex 4.3.5, Switzerland) [37].

2.3.3. Chemical characterization

The surface composition of the samples was characterized by Raman microscopy. An inVia™ Qontor® confocal Raman microscope (Renishaw, UK) was used, coupled to a specially adapted DM2700 M microscope (Leica, Germany). Spectra were acquired with an objective of 100 × , 10 accumulations of 10 s each, a laser with a nominal 100 mW output power, wavelength of 532 nm and a grating of 2400 lines mm⁻¹. For each measurement, three random measurements were taken per sample.

Additionally, the elemental composition analysis of the surfaces and cross sections were carried out using energy-dispersive X-ray spectroscopy (EDX) with an INCAPentaFETx3 detector (Oxford Instruments, UK). Before quantification, the electron beam was optimized at 20 keV using a cobalt reference standard. To determine the average surface atomic concentration, measurements were taken in at least three randomly selected areas on each sample. Additionally, line scans were conducted on cross-sectional samples to assess fluoride penetration depth. EDX data was smoothed using a Savitzky-Golay filter to remove the high frequency noise (20 coefficients, second order polynomial). Prior to measurements, samples were coated with carbon as described before.

X-ray photoelectron spectroscopy (XPS) was employed to analyze the surface chemical composition of the samples. The spectra were acquired in a SPECS system using an XR50 Al Kα line of 1486.6 eV anode source operating at 80 W, a MCD-9 detector and a PHOIBOS 150 EP hemispherical energy analyzer (SPECS Surface Nano Analysis GmbH, Germany). Survey spectra were acquired at a pass energy of 30 eV with an energy step of 1 eV. Narrow spectra were recorded at a pass energy of 20 eV and with step size of 0.1 eV, placed at 54° respect to analyzer axis and calibrated by the 3d 5/2 line of Ag with a full width at half maximum of 1.211 eV. A flood gun was used to compensate charging effects at 10 μA and 1 eV. All analysis was conducted under a vacuum pressure of less than 7.5 × 10⁻⁹ mbar. Binding energies were referenced against the C 1s signal set at 284.8 eV of adventitious carbon to correct charging effects. For each working condition, two samples were analyzed, with measurements taken at three different points per sample.

2.3.4. Contact angle

The sessile drop method was employed to evaluate the surface hydrophilicity using a Contact Angle System (OCA15 plus, Dataphysics, Germany). A droplet volume of 3 μL was dispensed at a rate of 1 μL min⁻¹. Ultrapure water (Millipore Milli-Q, Merck Millipore Corporation, USA) and diiodomethane (Sigma-Aldrich, Spain) were used as probe liquids. For each series, contact angle measurements were performed on ten independent samples for each condition at room temperature. Data were analyzed with SCA 20 software (Dataphysics), and the surface free energy was calculated using the Owens–Wendt method [38].

2.3.5. Mechanical properties

Vickers microhardness was measured on the surface of the nanostructured discs using a digital microhardness tester (Akashi MVK-H0, Japan). Flat surfaces were used as controls. A diamond indenter with a square pyramidal geometry and a vertex angle of 136° was applied under a load of 200 gf (HV0.2) for 20 s. The distance between adjacent indentations was maintained at a minimum of 300 μm to avoid interference effects. Following microindentation, the tested areas were examined by SEM to characterize the surface microstructure. For quantitative analysis, the two diagonals of 15 individual indentations were measured.

For compressive mechanical testing, cylindrical samples (∼5 mm in diameter and ∼7 mm in height) were prepared from α-TCP paste mixed with deionized water at a liquid-to-powder ratio (L/P) of 0.35. The paste was cast into plastic molds and incubated for 5 h at 37 °C under 100 % relative humidity to obtain solid cylinders. These cylinders were subsequently subjected to either autoclave or biomimetic treatments, as described in Section 2.2.2. Twelve wet samples per condition were tested longitudinally, and the Young's modulus and the ultimate compressive strength (UCS) were determined from compression tests performed using a universal testing machine (Bionix 858 Test System, MTS, USA) at a crosshead speed of 1 mm/min until fracture.

2.4. Biological characterization of the nanostructured surfaces

2.4.1. Antimicrobial properties of the CDHA nanotopographies

Bacterial assays were conducted using two strains: Gram-negative P. aeruginosa (ATCC 27853) and Gram-positive S. aureus (ATCC 25923). P. aeruginosa was cultured and maintained in LB, while S. aureus was grown in BHI.

To ensure mid-logarithmic phase culture, P. aeruginosa was first incubated for 8 h at 37 °C, then serially diluted (10⁻⁶ to 10⁻¹⁰) and incubated overnight. The resulting exponential-phase cultures were adjusted to an optical density (OD₆₀₀) of 0.1 and further diluted 1:10 for experimental use. For S. aureus, an overnight culture at 37 °C was prepared, from which a fresh bacterial suspension was adjusted to an OD₆₀₀ of 0.1 and incubated for 2 h to promote growth. This suspension was then standardized to an OD₆₀₀ of 0.1 ± 0.01 for the assays. In each experiment, flat controls containing fluoride, Flat and Ti samples were included as controls. All assays were performed under static conditions, with three biological replicates conducted for each experimental condition.

2.4.1.1. Bacteria interaction with the CDHA nanotopographies

Antibacterial effect was assessed with the LIVE/DEAD BacLight bacterial viability kit. Samples were sterilized by immersion in 70 % ethanol for 30 min, 3x rinsed with PBS, and subsequently immersed in 300 μL of bacterial suspension for 6 h at 37 °C (OD₆₀₀ of 0.1/10 for P. aeruginosa and 0.1 for S. aureus) in 48-well plates. Next, the medium was discarded, discs were washed three times with PBS and adherent bacteria were fixed with a solution of glutaraldehyde (2.5 % in PBS) for 30 min at room temperature. Upon fixation, the samples were 3x washed with PBS and stained by incubation with 300 μL of the dye-solution (2 μL of the mixture 67 mM SYTO9/18.3 mM propidium iodide in 1 mL of PBS) for 15 min. Dyed samples were analyzed by confocal laser scanning microscopy (CLSM, Zeiss LSM 800) equipped with an Ar laser, with the excitation/emission wavelengths of 561/550–650 nm and 488/410–546 nm and with a 63 × oil immersion objective. Three images for each condition were taken and analyzed using ZEISS software (Carl Zeiss, Germany). The ratio of dead bacteria was calculated by dividing the number of dead bacteria (red fluorescence and co-localized fluorescence) by the total number of bacteria (red and green fluorescence) and multiplying by 100.

For evaluation of bacterial metabolism, after sterilization of the samples, the substrates were incubated with 300 μL of bacterial suspension for 6 h at 37 °C, as previously explained. Following incubation, the medium was removed and the samples were 3x rinsed with PBS. Subsequently, 75 μL of fresh medium and 75 μL of BacTiter-Glo Reagent were added to each disc and incubated in the dark at room temperature for 5 min. Finally, 100 μL from each sample was transferred to an opaque white 96-well plate and luminescence was measured using a multimode microplate reader (Infinite® 200 PRO, TECAN, Switzerland). In order to normalize the BacTiter-Glo measurements versus the number of bacteria adhered to the surface of the discs, the supernatant was removed and adherent bacteria were fixed with a solution of glutaraldehyde (2.5 % in PBS) for 20 min at room temperature. Upon fixation, the samples were 3x washed with PBS and stained by incubation with Cellbrite 555 fix (1 μL of Cellbrite 55 fix in 1 mL of PBS) for 15 min. After rinsing 3x in PBS, dyed samples were analyzed by CLSM (Zeiss LSM 800) with the excitation/emission wavelengths of 561/555–650 and with a 63 × oil immersion objective. Three images were taken for each condition and analyzed using Fiji/ImageJ software (NIH) [36]. Final results were presented as the ratio of BacTiterGlo luminescence to the bacteria-covered area, and subsequently normalized to the Ti control (set as 100 relative luminescence units (RLU) per μm²).

2.4.1.2. Bacterial morphology

After bacterial incubation in the samples for 6 h, discs were washed 3x with PBS and bacteria were dehydrated through immersion in increasing concentrations of ethanol (10 % for 15 min thrice, 30 % for 15 min, 50 % for 15 min, 70 % for 15 min, 90 % for 15 min, 96 % for 15 min and 100 % for 10 min thrice). Then, the specimens underwent a graded transition to HMDS using mixtures of ethanol (100 %) at ethanol:HDMS radios of 2:1, 1:1 and 1:2 (v/v) for 15 min each. Finally, the samples were immersed in HDMS for 15 min and dried overnight at room temperature. Specimens were coated with 6 nm of iridium (high vacuum sputtering Leica EM ACE600, Leica, Germany) for examination under TESCAN AMBER FIB-SEM (Tescan, Czech Republic) equipment. For each sample, five images were taken at a working distance of 4 mm and a potential of 2 kV.

2.4.2. Cell experiments

The SaOS-2 cell line (HTB-85) was cultured in McCoy's 5A medium supplemented with 15 % (v/v) FBS, and 1 % p/s at 37 °C in a humidified incubator with 5 % (v/v) CO₂. The mouse macrophage RAW 264.7 cell line was cultured in DMEM supplemented with 1 mM sodium pyruvate, 10 % (v/v) FBS and 1 % (v/v) p/s at 37 °C in a humidified incubator with a 5 % CO₂ level. The medium was refreshed every two days and passages between 33 and 37 (for SaOS-2) and between 20 and 30 (for RAW 264.7 cells) were used. Prior to reaching confluence, cells were 3x washed with PBS and detached from the culture flask either by incubation with TrypLE (for SaOS-2 cells) or by using a cell scraper (for RAW 264.7 cells). When using TrypLE, after 5 min incubation time at 37 °C, the cells were centrifuged at 300g for 5 min, and resuspended. Appropriate aliquots of the cell suspension were transferred to a new culture flask for further cultivation. The discs were sterilized as explained in section 2.4.1.1 and placed in a 12-well plate. The discs were preconditioned by incubation with complete McCoy's 5A medium (4 mL, for SaOS-2) or DMEM (4 mL, for RAW 264.7) in a humidified atmosphere of 5 % CO₂ at 37 °C for 24 h. After preconditioning, the medium was removed, and SaOS-2 cells were seeded at 300 cells mm⁻² onto the discs. RAW264.7 seeding densities were adjusted according to the incubation period to ensure maximum surface coverage by the time of RNA or protein extraction. The same number of cells was seeded on the different substrates because the nanoscale features present on the surfaces do not substantially modify the effective surface area available for cell growth. As all samples had identical macroscopic dimensions (disc diameter), the total available area for cell attachment was considered equivalent across conditions. Ti discs were used as controls.

2.4.2.1. Cell proliferation and morphology

The proliferation of SaOS-2 cells cultured on the different surfaces was evaluated using the Cytotoxicity Detection Kit LDH. After 4 h, and at 7, 14, and 21 days of incubation in complete medium, samples were 3x washed in PBS and cells were lysed with 200 μL well⁻¹ of M-PER®. The released LDH, which correlates with the number of proliferating cells, was quantified spectrophotometrically measured at 490 nm with a microplate reader (Infinite® 200 PRO, TECAN, Switzerland). A calibration curve was prepared with a known number of cells to establish a correlation between absorbance and number of viable cells. Ti samples were included as positive controls. Each condition was evaluated by triplicate.

Nanotopography samples with cells were incubated at 37 °C and 5 % CO₂ in complete medium. After different time incubations, discs were 3x rinsed with PBS to remove loosely adherent cells. Then, samples were fixed with a solution of PFA (4 % in PBS) for 30 min at room temperature and 3x washed in PBS-Gly. Cells were permeabilized with 0.1 % Triton X-100 in PBS for 20 min at room temperature and 3x washed with PBS-Gly. The samples were stained with Alexa Fluor™ 546 phalloidin (1:300 in PBS) for 1 h in the dark. After 3x rinses in PBS-Gly, the discs were incubated for 2 min in the dark at room temperature with a solution of DAPI (1:1000, in PBS). Following three additional PBS-Gly washes, the nanotopography surfaces were imaged using CLSM (Zeiss LSM 800, Carl Zeiss, Germany) equipped with a 40 × oil immersion objective. Excitation/emission wavelengths were set to 405/440–480 nm for DAPI and 561/555–650 nm for Alexa Fluor™ 546 phalloidin.

2.4.2.2. Alkaline phosphatase activity

The alkaline phosphatase (ALP) activity of SaOS-2 cells was assessed over a period of 21 days, with Ti serving as the control. SaOS-2 cells were seeded onto the samples, lysed and following the same protocol used for the cell proliferation assay (refer to Section 2.4.2.1). Cell lysates were incubated with a p-nitrophenol phosphate (pNPP) solution at room temperature in the dark for 10 min, using the SensoLyte® pNPP Alkaline Phosphatase Assay Kit. The absorbance of the resulting p-nitrophenol was measured at 405 nm with a multimodal plate reader (TECAN). The total p-nitrophenol concentration was determined using a calibration curve, and the enzymatic ALP activity was expressed as ng of the reaction product (p-nitrophenol) per minute (30 min total). Total protein content was measured using BCA protein assay kit according to the manufacturer's protocol, and ALP activity values were normalized to the total protein content.

2.4.2.3. Gene expression

The RUNX2, ALPL, and SPP1 osteogenic-related genes and TNF, IL-1β and IL6 inflammatory-related genes were considered. Primer sequences are shown in Table S1. Total mRNA was extracted from the samples at different time points using the RNeasy Mini Kit according to the manufacturer's instructions. RNA purity and concentration were determined by measuring absorbance at 260 nm using a micro-spectrophotometer Nano-400A (Allsheng, China). The purified RNA was stored at −80 °C until further use. For the RT-qPCR, 100 ng of RNA was reverse transcribed into cDNA and subsequentially amplified using the SuperScript™ III Platinum™ SYBR™ Green One-Step Kit, following the manufacturer's protocol. The amplification protocol consisted of an initial reverse transcription step at 50 °C for 10 min, polymerase activation at 95 °C for 1 min, followed by 40 cycles of denaturation at 95 °C for 15 s and annealing/extension at 55 °C for 30 s. Gene expression was assessed with an AriaMx Real-Time PCR System (Agilent, CA, USA) and normalized to the β-actin housekeeping gene for SaOS-2 cells and GAPDH housekeeping for RAW 264.7 cells. Relative expression levels were calculated using the Livak's (2⁻ΔΔCq) method, with cells cultured directly on titanium at day 1 serving as the reference condition.

2.4.2.4. Inflammatory antibody array

To evaluate the expression of inflammation-related proteins in RAW 264.7 cells, an inflammatory antibody array was used. RAW 264.7 cells were seeded on the treated discs for 7 days. For the last 3 days, cells were cultured in absence of FBS to prevent the appearance of background signal in the array and the supernatant was stored at −80 °C. The samples were then 3x washed with PBS and the cells were lysed with 200 μL of MPER reagent. Subsequently, both the conditioned medium and the cell lysates were mixed and concentrated using Centrifugal Filter Units (3 kDa), in a total of 1 mL. Protein concentration in the resulting volumes was quantified by BCA assay and adjusted to 1 mg total protein per 1 mL. The protein quantification was then performed using the mouse inflammation antibody array following the manufacturer's protocol and the signal was detected using a Western blot developer Syngene™ GeneGnome XRQ (Fisher Scientific). RAW 264.7 cells seeded in titanium were used as control. The signal was quantified using Fiji/ImageJ software, and the values were relativized to the control.

2.4.3. In vitro co-culture evaluation

After sample sterilization, P. aeruginosa and SaOS-2 cells were co-cultured, varying the seeding order to simulate two distinct clinical scenarios: bacteria seeded first to model a pre-implantation infection, and cells seeded first to represent a post-implantation infection. For all experiments, discs were sterilized as described in section 2.4.1.1 and preconditioned by incubation with complete McCoy's 5A medium (4 mL) in a humidified atmosphere of 5 % CO₂ at 37 °C for 24 h.

2.4.3.1. Pre-implantation infection protocol

P. aeruginosa was incubated until it reached the mid-exponential phase, and then it was diluted to OD₆₀₀ of 0.1 in antibiotic-free cell medium (McCoy's media supplemented with 1 % FBS and 20 mM HEPES buffer). Samples were then immersed in 300 μL of bacterial solution (further diluted 1:10 just prior usage) and incubated for 6 h at 37 °C. Following incubation, samples were 3x washed with PBS. SaOS-2 cells (1200 cells mm⁻²) were then seeded in antibiotic-free cell medium and incubated for 24 h at 37 °C in a humidified incubator with 5 % CO₂ (v/v).

2.4.3.2. Post-implantation infection protocol

SaOS-2 cells (1200 cells mm⁻²) were seeded onto the modified CaP discs and incubated for 24 h at 37 °C in a humidified incubator with 5 % CO₂ (v/v). After incubation time, samples were 3x washed with PBS. Bacterial solution (300 μL) at to OD₆₀₀ of 0.1/10 in antibiotic-free cell medium was added and incubated for 6 h at 37 °C in a humidified incubator with 5 % CO₂ (v/v).

In the co-cultures, cell morphology was evaluated by fluorescence staining as it has been described in section 2.4.2.1. CLSM images were captured (Zeiss LSM 800) and analyzed with Fiji/ImageJ software [36]. Early bacterial colonization was assessed using a LIVE/DEAD BacLight bacterial viability kit as has been described previously in section 2.4.1.1. Three samples were studied for each condition.

2.5. Statistical analysis

For nanostructures length and distances, one-way ANOVA followed by multiple comparison correlation and Tukey's post hoc test was used. Non-parametric Kruskal-Wallis and Mann–Whitney U test were used to analyze significant differences in Raman spectroscopy, mechanical measurements and biological tests (i.e. cell proliferation assays, bacterial live/dead staining, BacTiterGlo and gene expression analyses). Significance level was set at a P value of ∗p ≤ 0.05, ∗∗p ≤ 0.01, ∗∗∗p ≤ 0.001 and ∗∗∗∗p ≤ 0.0001. All data were analyzed in SPSS software (IBM, NY, USA).

3. Results and discussion

3.1. Synthesis of F-doped nanotopographies and characterization

Bone graft failure due to bacterial infection and subsequent biofilm formation remains a critical challenge in tissue engineering [39,40]. In response, extensive research has been devoted to developing surface modification strategies to prevent microbial colonization [[41], [42], [43]]. Among these, nanotopographic surface modifications have emerged as an encouraging approach [44,45], offering a purely physical mode of antibacterial action that avoids the drawbacks associated to conventional antimicrobial agents. In this study, we build upon this strategy by integrating nanotopographical features with ionic cues through fluoride incorporation. This combined approach aims to explore potential synergistic effects between topographical and chemical mechanisms to enhance overall antibacterial performance.

3.1.1. Morphological analysis

Nanostructured CDHA samples were developed using a previously established method [46,47]. This process involves the hydrolysis of α-TCP, triggered by localized calcium and phosphate supersaturation, followed by precipitation of CDHA crystals [48,49]. In this study, we incorporated fluoride ions in the soaking solution, aiming to tailor both structural and chemical properties of the material. The hydroxyapatite structure is very prone to incorporating ionic substitutions within its crystal lattice, which can significantly affect crystal nucleation and growth [[24], [25], [26]]. For example, the incorporation of carbonate ions, very common in biological apatite, leads to smaller less crystalline particles, with smaller aspect ratio [50,51]. Similarly, the zinc ion inhibits crystal growth by reducing crystallite size and thermodynamic stability [52,53]. In contrast, fluoride substitution for hydroxyl groups has been reported to promote the formation of more elongated and well-ordered apatite crystals [25,27,30]. We hypothesized that the presence of fluoride ions in the soaking solution would promote the precipitation of fluoride-doped hydroxyapatite, thereby influencing surface nanotopography. Simultaneously, the presence of fluoride on the surface was expected to further enhance the antibacterial properties.

The morphology of the nanotopographies is presented in Fig. 2a, which displays the cross-sectional SEM images of the CDHA surfaces, revealing the formation of nanoneedles under all experimental conditions. Compacted biomimetic CDHA was used as a Flat control. In the absence of fluoride, the topography resulted in a dense array of needles growing perpendicular to the disc surface. The length of these nanoneedles was notably longer using autoclave conditions compared to those formed under biomimetic conditions (∼1.740 μm and ∼430 nm for 0F_AC and 0F_BM, respectively). The morphological characteristics of these nanostructures were consistent with those reported by Iglesias-Fernandez et al. who observed crystal heights of ∼640 nm and ∼1.5 μm for biomimetic and autoclave treatment, respectively, aligning well with our findings (Fig. 2b) [23]. The observed differences in crystal morphology arise from the distinct kinetics of the two synthesis routes: while the lower-temperature biomimetic approach produced finer CDHA crystals, the higher temperature and pressure of the autoclave method resulted in more pronounced crystal growth.

During CDHA formation in the presence of fluoride, a clear trend was observed under autoclave treatment. We observed a decrease in needle length with increasing fluoride concentration. This finding was contrary to our initial expectations, as the literature suggests that fluoride generally promotes preferential c-axis growth, resulting in more elongated hydroxyapatite crystals [27,30,54]. This unexpected divergence may be attributed to the specific synthesis parameters and accelerated reaction kinetics of the autoclave methodology. In contrast, no such fluoride-dependent trend in needle length was observed in the biomimetic treatment, where crystal dimensions remained similar across all fluoride concentrations. Based on the morphological characteristics, six distinct nanotopographies with varying needle lengths were identified and selected for subsequent experiments. Fluoride concentrations of 0, 0.5 and 10 mM were chosen based on significant differences in needle length for both autoclave and biomimetic treatment.

To quantify needle density and inter-needle spacing resulting from different treatment conditions, SEM images of the disc surfaces were acquired (Fig. 3a). SEM analysis revealed notable variations in needle density among the samples. Under BM conditions, a densely packed needle network was observed, with densities of 184.2, 224.6 and 182.3 needles μm⁻² for 0F_BM, 05F_BM and 10F_BM respectively (Fig. 3b). In contrast, the 0F_AC discs exhibited densely clustered needle arrangements, with large inter-cluster cavities. For quantification, each cluster was considered as a single needle, resulting in a significantly reduced density of 4.5 needles μm⁻². However, within the AC group, increasing fluoride content was associated with a progressive increase in needle density, likely due to a reduction in cluster formation. Specifically, the 05F_AC and 10F_AC samples exhibited 45.8 and 121.7 needles μm⁻², respectively.

The inter-needle spacing, inversely correlated with needle density, was subsequently quantified (Fig. 3c). The 0F_AC substrate displayed a broad inter-needle spacing distribution (from ∼250 to ∼1500 nm), while the 05F_AC and 10F_AC samples exhibited narrower distributions (from ∼100 to ∼500 nm). Notably, increasing fluoride content in the AC samples resulted in a marked decrease in inter-needle spacing (mean spacing of 973.2, 236.4 and 169.2 nm for 0F_AC, 05F_AC, and 10F_AC, respectively). In contrast, the BM groups showed consistent inter-needle spacing distributions and mean values (∼140 nm) across all fluoride concentrations.

A prevailing hypothesis attributes the bactericidal effects of nanostructured surfaces primarily to mechanical rupture of the bacterial cell wall. This rupture is thought to arise from mechanical strain induced by contact between the bacterial cell wall and the nanoneedles, generating tensile stress between adhered and suspended regions of the cell [55]. Consequently, inter-feature spacing emerges as a critical parameter in the design of antibacterial topographies. Previous studies have shown that naturally occurring antibacterial surfaces typically exhibit feature spacings below 500 nm [56]. In our study, nearly all the fabricated topographies fall within this effective range, except for the 0F_AC sample, which exceeds this value. However, some natural antibacterial structures, such as gecko spinules, and synthetic nanotopographies maintain antibacterial efficacy despite exhibiting inter-structural spacing above this threshold [56]. Regarding needle height, most natural bactericidal topographies are in the range between 100 and 900 nm, but several antimicrobial artificial nanotopographies lie outside this range [13]. Therefore, both the needle height and inter-needle spacing dimensions of our selected surfaces align well with the morphological criteria reported for effective antibacterial nanotopographies.

3.1.2. Phase characterization

Fluoride ions (F⁻) can substitute hydroxyl ions (OH⁻) at the anionic sites of the apatite lattice, owing to their similar ionic radii (1.32 Å for fluoride vs. 1.68 Å for hydroxyl ions) [57]. This allows for the formation of a continuous solid solution across the full compositional range between hydroxyapatite (Ca₁₀(PO₄)₆(OH)₂) and fluorapatite (Ca₁₀(PO₄)₆F₂). Partial substitution yields intermediate compositions described by the general formula Ca₁₀(PO₄)₆F_x(OH)_2-x, for stoichiometric hydroxyapatite and Ca₉(PO₄)₅(HPO₄)F_x(OH)_1-x for CDHA with a Ca/P 1.5.

To elucidate the effect of the different hydrolysis treatments and fluoride doping on the phase composition of the samples, XRD and Raman spectroscopy were employed (Fig. 4). XRD analysis (Fig. 4ai and 4aii) revealed that the samples synthesized under BM conditions consisted predominantly of CDHA, with a small fraction of residual α-TCP and β-TCP. The latter phase was already present in the starting α-TCP, likely stabilized during the quenching process. Samples treated by autoclave exhibited a higher β-TCP content. The allotropic α-to-β allotropic transformation of TCP has been previously reported by other authors during the hydrolysis of α-TCP at elevated temperatures and pressures [20,23,58]. Notably, fluoride incorporation exerted a stabilizing effect on the apatitic phase, leading to an increased fraction of CDHA and a corresponding reduction of β-TCP content after AC treatment. The autoclave treatment also resulted in higher crystallinity, as indicated by the sharper peaks compared to BM substrates. Grazing incidence XRD (GIXRD) analysis (Fig. 4aiii) revealed the shift of the peaks at 31.7° and 32.8°, attributed to the (211) and (300) crystal planes, to higher angles with increasing fluoride concentration, consistent with the substitution of hydroxyl groups by fluoride ions within the hydroxyapatite lattice [[59], [60], [61]].

Raman spectroscopy confirmed the presence of the typical vibrational bands for the PO₄³⁻ and HPO₄²⁻ groups of CDHA in all samples (Fig. 4bi) [[62], [63], [64]]. The main PO₄³⁻ vibrational modes identified included the doubly degenerate bending (υ₂PO₄) at 430 and 446 cm⁻¹, the triply degenerate bending (υ₄PO₄) at 580, 590 and 605 cm⁻¹, the symmetric stretching (υ₁PO₄) around ∼960 cm⁻¹ and the triply degenerate asymmetric stretching (υ₃PO₄) at 1030, 1047 and 1076 cm⁻¹. Additionally, the HPO₄²⁻ vibration bands associated to the P-OH stretching were also found at 875 and 1002 cm⁻¹. The detection of these HPO₄²⁻ groups specifically confirms the non-stoichiometric nature of the hydroxyapatite, thereby demonstrating its successful formation [64]. Notably, the Raman shift at ∼946 cm⁻¹, attributable to β-TCP was not detected, indicating that this phase was not exposed on the surface of the discs [65]. Moreover, fluoride substitution for hydroxyl groups in the CDHA structure was also confirmed by Raman spectroscopy. Key spectral evidence included a shift in the PO₄³⁻ stretching vibration band (∼960 cm⁻¹) towards higher wavenumbers (Fig. 4bii) and a decrease in the OH⁻ vibration intensity (∼3570 cm⁻¹), notably, with its near absence in the 10F_BM sample (Fig. 4biii). These findings demonstrate successful fluoride incorporation, via hydroxyl group substitution, in turn influencing unit cell structure and phosphate group vibrations [61,66,67]. To further quantify fluoride incorporation, the phosphate-to-hydroxyl ratio was analyzed, as this parameter reflects the extent of hydroxyl substitution by fluoride within the CDHA structure. As shown in Fig. 4biv, the ratio increased with rising fluoride concentrations during nanostructure synthesis, through the trend differed between the BM and AC samples. In the AC group, changes in the bands associated with the substitution of OH⁻ by fluoride were only evident at the highest fluoride concentration (10F_AC). In contrast, the BM samples exhibited more pronounced spectral modifications, suggesting greater fluoride incorporation into the CDHA lattice, with complete substitution of OH⁻ for fluoride ions in the 10F_BM group.

3.1.3. Elemental composition

The surface composition of all nanostructured surfaces was determined using EDX (Fig. 5a). Fluoride incorporation led to a significant increase in fluoride content and a corresponding reduction in oxygen levels in the fluoride-treated samples compared to the controls (0F_AC, 0F_BM, and Flat). In both AC and BM treatments, the fluoride content increased gradually with its concentration in the soaking solution. The fluoride content of stoichiometric fluorapatite corresponds to 4.76 at%, whereas calcium deficient fluorapatite with a Ca/P ratio of 1.5 contains 2.44 at% of fluoride. The combination of the EDX data together with the progressively disappearing OH⁻ band in the Raman spectra suggests that fluoride exists in two different states on the sample. While some fluoride was incorporated directly into the apatitic crystal structure, an additional amount is likely adsorbed on the surface or physically trapped between the crystals. For instance, the amount of fluoride detected in the BM samples significantly exceeded the theoretical maximum that can be incorporated into the apatite structure, even assuming complete substitution of hydroxyl groups by fluoride ions. In contrast, the surface fluoride content in the autoclaved samples was lower. In the case of the 10F_AC sample, part of the fluoride appeared to be structurally incorporated via partial OH⁻ substitution, whereas in the 05F_AC sample, the Raman results suggest that the fluoride was entirely surface-bound and not incorporated into the apatitic lattice.

In the same line, fluoride incorporation did not significantly affect the Ca/P ratio in the AC samples, which remained consistently around 1.5 across all samples. In contrast, in BM samples, fluoride incorporation was accompanied by an increase in the Ca/P ratio, in agreement with the results reported by Rodriguez-Lorenzo et al., for solid solutions of hydroxy-fluorapatites [68].

Moreover, cross-sectional analysis of the discs was conducted to confirm fluoride incorporation and assess the depth of doping (Fig. 5b). EDX revealed fluoride at the surface of 10F_AC, 05F_BM and 10F_BM samples, with a gradual decrease in concentration along the depth. The penetration depth varied from approximately 15 μm in 10F_AC to 20 μm in 05F_BM, reaching up to 50 μm in 10F_BM. The greater depth observed in BM samples is attributed to the longer incubation times of the BM process, which allows fluoride ions to diffuse more extensively into the material, consistent with Fick's second law of diffusion [69]. Taken together, our analysis demonstrated the successful incorporation of fluoride into the CaP discs using the applied methodology and the superior efficacy of the BM treatment compared to the AC approach for fluoride incorporation.

To further clarify the specific contribution of fluoride to the antibacterial effect independent of the surface nanotopography, control samples were prepared by re-pressing the fluoride-doped materials to eliminate the nanostructure. As shown in Table S2, the fluoride atomic content of the compacted samples was consistent with that of their nanostructured counterparts, thereby enabling a direct and reliable comparison of the antibacterial effects due to the chemical composition.

XPS measurements were performed to investigate the chemical composition and bonding states of the different samples (Fig. 6). The survey spectra (Fig. 6a) confirmed the presence of the expected elements (Ca, P, O, C, and F). A progressive increase in fluorine content was detected with increasing fluoride concentration in both AC and BM series, although to different extents, following the same trends observed by EDX (Fig. 5a). The discrepancies in the respective values can be associated with the greater penetration depth of EDX compared to the nanometer-scale probing depth of XPS.

High-resolution spectra and deconvolution of the F 1s region (Fig. 6b) revealed two distinct components. A first peak located at ∼684.2 eV can be assigned to fluoride ions substituting hydroxyl groups in the HAp lattice, characteristic of fluoridated HAp (FHAp), in agreement with previous reports [70,71]. A second peak observed at ∼686.6 eV is attributed to fluoride ions bound to calcium at the surface, consistent with the formation of Ca–F bonds [70,71]. Both contributions were present in all fluoridated samples, with their relative intensity depending on fluoride concentration and synthesis conditions, as shown in Fig. 6bii. For instance, 05F_AC showed 15 % of the component corresponding to fluoride bound to calcium on the surface, and 85 % to fluoride substitution in the apatitic lattice, while in 10F_BM lattice substitution represented 93 % and surface bonding only 7 %. These findings confirm that fluoride is incorporated both as lattice-substituted F⁻ within HAp and as surface Ca–F species. The coexistence of these bonding environments highlights the dual role of fluoride in modifying both bulk and surface chemistry, which is expected to influence nanocrystal growth, and also surface energy, wettability, and ultimately biological responses.

3.1.4. Contact angle analysis

The wettability and surface energy (SE) parameters of the samples are summarized in Fig. 7a. All nanostructured surfaces showed lower CA than the flat control, indicating increased hydrophilicity. AC samples exhibited the lowest CA values (5.91–8.57°), while BM samples presented slightly higher values (9.31–14.53°), compared to the 20.03° of the Flat control. The incorporation of fluoride resulted in a slight increase in the CA of the AC samples, approaching the values observed for the fluoride-doped BM samples. This is consistent with the reduction in nanopillar height induced by fluoride incorporation in the AC samples, and suggests that nanostructure, rather than the presence of fluoride itself, is the primary contributor to the change in CA. Both AC and BM treatments markedly increased total SE compared to the Flat control (73–74 mN/m vs. 20 mN/m), mainly due to an increase in the polar component (40–41 mN/m). The concurrent decrease in CA and increase in SE confirm that nanotopographies amplify surface wettability, in agreement with the Wenzel model [72]. The nanostructured surfaces demonstrated similar polar contributions, comparable also to the value obtained for the flat substrate, and similar affinity with water molecules through hydrogen bonding. Surfaces with higher polarity generally favor protein adsorption and cell adhesion while potentially limiting nonspecific bacterial attachment, but in this case the similarity among the different substrates are not expected to cause distinct behaviors in this respect.

3.1.5. Mechanical properties

The mechanical response of the nanostructured surfaces was assessed by microhardness and compression testing, and the results are presented in Fig. 7b and c. Vickers microhardness measurements revealed no statistically significant differences among the tested conditions, including the Flat control. All samples exhibited values within the range of 80–100 HV0.2 (Fig. 7biii). These values are slightly higher than those typically reported for natural bone (40–60 HV0.2) yet remain within an acceptable range for mechanical compatibility with biological tissue [73].

The compressive properties of the pristine 0AC and 0BM samples were similar in terms of both elastic modulus and UCS. The incorporation of fluoride produced a slight reduction in the elastic modulus for the BM samples, and some variations in UCS were observed in both groups. However, in all cases, the measured values remained above those typically reported for cortical bone [74,75] and for non-sintered calcium phosphate materials, [[76], [77], [78]], making them suitable for bone grafting applications.

3.2. Antibacterial activity

3.2.1. Early bacterial colonization

Following the successful synthesis and characterization of the nanotopographical modifications on CaP surfaces, their antimicrobial properties were evaluated against P. aeruginosa (Gram-negative) and S. aureus (Gram-positive) strains. These strains were selected due to their prevalence in chronic wound infections and established antimicrobial resistance [79].

Initial bacterial colonization of P. aeruginosa was assessed using a live/dead assay (Fig. 8ai and Fig. S1). Nanostructured CDHA samples demonstrated significant inhibitory effects on bacterial colonization compared with the Flat CDHA but no statistically significant differences were observed between the two nanostructures despite the differences in needle spacing and needle length. Notably, fluoride-doped samples exhibited the most pronounced inhibitory activity, with no statistically significant differences observed among them. However, an increase in the antimicrobial properties was observed when comparing fluoride-doped and non-doped samples, as statistically significant differences were detected. To isolate the contribution of fluoride from that of nanotopography, fluoride-containing compacted samples (C05F_AC, C10F_AC, C05F_BM, and C10F_BM) were analyzed. These samples retained fluoride content but lacked surface nanoneedles due to the compaction process. The results revealed no significant antibacterial activity against P. aeruginosa in any of the compacted groups, indicating that fluoride alone, at the tested concentrations, did not exert an independent antibacterial effect.

The absence of antibacterial activity from the solitary ion suggests the requirement of an alternative mechanism; therefore, we must consider the physical contribution of the surface. The antibacterial performance of nanostructures has been shown to be dependent on their geometry. While sharp, high-aspect-ratio features effectively kill bacteria by inducing high mechanical stress, larger structures tend to be less effective [13,[80], [81], [82]]. However, within the ranges covered in the present study, we did not observe a higher antimicrobial efficacy for the longer nanoneedles of the 0F_AC surfaces, compared to the shorter nanoneedles of the 0F_BM surfaces (Fig. S2). The combined results from multiple experiments suggested a synergistic antibacterial effect. First, control samples containing fluoride but lacking nanotopography (i.e., the compacted discs) exhibited no antibacterial activity. Second, MIC and agar diffusion tests showed that the amount of fluoride in the discs was too low to exert a direct antibacterial effect and that the fluoride released into the medium was insufficient to inhibit bacterial growth (Fig. S3). Finally, the BacTiter-Glo assay confirmed these findings, revealing a significant reduction in metabolism across all nanostructured samples, an effect most pronounced on the fluoride-containing surfaces (Fig. 8aii). This trend was consistent for both the BM and AC series, confirming the robustness of the synergy across different nanotopographies, including the varied surface topographies of the AC samples at different fluoride concentrations. Taken together, these results demonstrate that the enhanced antibacterial performance does not result from a simple additive combination of topographical and ion-doping effects, but rather reflects a synergistic interplay between the surface nanotopography and the incorporated fluoride.

Nanostructured surfaces can physically deform, stretch or puncture bacterial membranes [13,83,84], compromising membrane integrity and interfering with osmotic regulation [85]. The resulting damaged or stressed membrane likely facilitates greater fluoride uptake, inhibiting key metabolic enzymes such as enolase, thereby reducing ATP production and acid generation, intensifying intracellular acidification and proton motive force [34]. Thus, a possible mechanism underlying this synergy might be that although the total ion content is similar in both compacted and nanostructured specimens, the nanotopographies expose fluoride at the surface in a way that increases its accessibility and interaction with bacteria, thereby enhancing its effect.

To evaluate the broader applicability and robustness of the antibacterial nanotopographies, their efficacy was further tested against S. aureus, a clinically relevant Gram-positive pathogen known for its thick peptidoglycan layer, higher resistance to mechanical disruption, and its prominent role in orthopedic implant-associated infections. Live/dead staining (Fig. 8bi and Fig. S1) revealed no significant bactericidal effect of the tested nanostructured surfaces against S. aureus, indicating limited membrane disruption under these conditions. However, metabolic activity assays (Fig. 8bii) demonstrated statistically significant reductions in bacterial viability in the nanostructured samples. Specifically, 05F_AC and 10F_AC exhibited significant reductions in metabolic activity compared to 0F_AC, while 10F_BM showed a substantial decrease compared to 0F_BM. As for P. aeruginosa, the fluoride-containing compacted samples exhibited no detectable antibacterial effect, as confirmed both by the live/dead and in the BacTiter-Glo assays.

The observed discrepancies between the live/dead and metabolic activity assays for S. aureus, as well as its differential response compared to P. aeruginosa, may be explained by the different sensitivities of these bacterial strains to the nanotopographical features. Although the full mechanism is not yet completely understood and may involve other factors such as reactive oxygen species or steric hindrance, it is generally accepted that sharp nanoneedles physically deform or rupture the bacterial cell envelope [13,42,86]. Such mechanism appears to be effective against P. aeruginosa, as evidenced by the live/dead assay, which employs a red fluorescent dye to identify bacteria with compromised cell wall integrity. We hypothesize that S. aureus maintains its cell envelope integrity, thereby preventing the red dye from penetration. Conversely, the BacTiter-Glo assay, which measures intracellular ATP concentration and is independent of cell wall integrity, revealed a reduction in metabolic activity of S. aureus on the developed nanotopographies. While these findings suggest an alternative non-lytic mechanism of action against S. aureus, further investigations are warranted to elucidate the underlying processes. However, such mechanistic studies are beyond the scope of the present work.

From the compositional point of view, the material may affect bacterial viability not only through contact-based mechanisms but also via ion exchange, which may have chemical toxicity. To better understand if the inhibition or killing of bacteria was also influenced by the ionic concentration in solution, the ion exchange between the samples and the bacterial medium was investigated using ICP-MS (Fig. S4). For P. aeruginosa, supernatants from the Flat sample showed the highest calcium concentration. This elevated release can be attributed to the higher specific surface area (SSA) and higher porosity of the sample, which consists of compacted nanocrystals of CDHA, compared to the hydrolyzed α-TCP discs, thus enhancing its reactivity (Table S3). In contrast, when tested with S.aureus, the Flat sample exhibited a less pronounced ionic exchange, probably due to the higher calcium concentration in the BHI growth medium, which may have diminished the driving force for further ion release from the material. Overall, no drastic changes were observed in the Ca and P concentrations in the supernatants of the nanostructured samples, compared to the pristine LB and BHI broths or the chemically inert Ti control. This suggests that changes in ionic levels in solution are not the primary direct cause of the observed bacterial mortality, instead pointing towards the role of mechanical and direct-contact mechanisms of the fluoride-doped topographies.

Surface wettability also influences bacterial behavior. Although increased hydrophilicity and surface energy are often linked to improved biological performance, the similar CA and SE values among treated samples suggest that the differences in bacterial adhesion cannot be attributed solely to wettability. Instead, the antibacterial effect likely arises from the combined influence of nanoscale topography and surface chemistry, which create unfavorable conditions for bacterial attachment while maintaining compatibility with protein adsorption and cell adhesion.

3.2.2. Bacteria morphology

To examine bacterial morphology on the synthesized nanotopographies, SEM images were acquired. Fig. 8aiii illustrates the interaction between P. aeruginosa and the surface nanostructures, revealing direct contact between the nanoneedles and the bacterial cell wall. This contact is often associated with visible signs of cell wall stress or deformation. In contrast, SEM analysis of S. aureus (Fig. 8biii) showed no evidence of cell wall penetration or mechanical damage caused by the nanostructures, suggesting an alternative mechanism of bacterial inactivation.

3.3. Cell experiments

3.3.1. Cell proliferation and differentiation

Given the promising antibacterial properties of the engineered nanotopographical surfaces, it is crucial to ensure their biocompatibility with mammalian cells to support safe biomedical applications. To this end, the biocompatibility and osteogenic bioactivity of the synthesized nanostructures were assessed using human osteosarcoma-derived SaOS-2. It must be acknowledged that SaOS-2 cells are osteosarcoma-derived and may not fully replicate the behavior of primary osteoblasts. Their tumor origin is associated with altered adhesion dynamics and integrin expression compared to MSCs [87], as well as altered proliferation, differentiation, and metabolic activity relative to normal osteoblasts. Nevertheless, SaOS-2 cells are widely used in preliminary in vitro studies because of their reproducible growth, stable phenotype, and consistent expression of key osteoblastic markers. However, to better approximate physiological conditions, the response of primary cells should be investigated in future studies. Cell proliferation was monitored and ALP activity was quantified at various time points (4 h, 7, 14, and 21 days) to capture both short and long-term cellular responses. Fig. 9a presents the cell proliferation across all tested surfaces. After 4 h of incubation with cells, all sample groups presented comparable cell numbers, suggesting that initial adhesion was still in its early stages and was not negatively impacted by the material properties. At day 7, both Flat and Ti control groups exhibited increased cell numbers, while the nanostructured samples showed relatively constant levels. However, a significant increase in cell number was observed in all nanostructured samples by day 14, irrespective of fluoride incorporation. The Flat and Ti samples exhibited similar proliferation profiles until day 14. Beyond this point, Ti samples appeared to reach confluence, leading to a plateau in cell numbers between days 14 and 21. In contrast, the Flat sample continued to support cell proliferation through day 21. While nanostructured samples initially lagged behind the Flat controls, cell proliferation was not inhibited; rather, a sustained increase in cell numbers was observed over the duration of the study, suggesting good cytocompatibility despite early proliferation delays.

Fig. 9 — (a) Cell proliferation of SaOS-2 cells onto the different CaP discs for 4 h, 7, 14 and 21 days. (b) ALP activity of SaOS-2 cells cultured onto the CaP substrates for 4 h, 7, 14 and 21 days. The same letter indicates no statistically significant differences for the same group at different time points while the same number denotes no statistically significant differences for each time point among all samples. (p < 0.05).

Interestingly, the BM samples, despite significant differences in fluoride content, displayed comparable cell proliferation by day 21, likely due to their shared nanotopographical features. However, at day 14, the 05F_BM and 10F_BM groups showed higher cell numbers when compared to 0F_BM, potentially reflecting a fluoride-induced enhancement in cellular activity [88,89]. In contrast, the 0F_AC sample presented the most pronounced inhibition of cell proliferation, with cell numbers plateauing by day 14. This inhibitory effect is likely due to its distinct topographical architecture, characterized by a high aspect ratio and wide inter-needle spacing, which may limit the availability of adhesion sites and impair cell attachment and spreading, ultimately rendering the surface less conducive to cell growth.

Previous studies have shown that these materials with active ion exchange profiles can modulate cell behavior, including proliferation [90]. To evaluate whether such chemical interactions contributed to the observed cell responses, particularly proliferation inhibition on the 0F_AC's sample, ICP-MS analysis was performed. As shown in Fig. S5, Flat discs, exhibited increased reactivity and active calcium sequestering. This is likely a consequence of their higher specific surface area and porosity, as seen by nitrogen adsorption (Table S3). Notably, despite these ionic fluctuations, no detrimental effect on cell proliferation was observed, aligning with previous reports. In contrast, the 0F_AC sample, which displayed negligible ionic fluctuations compared to the rest of the samples, still exhibited cell proliferation inhibition. This suggests that the observed cytocompatibility issues are not driven by the ionic exchange. In the specific case of 0F_AC, the reduction in cell proliferation is most likely associated to the topographical features of the sample. Importantly, the proliferation assays confirmed that, with the exception of the 0F_AC sample, neither the nanostructure nor the incorporation of fluoride induced cytotoxic effects on SaOS-2 cells.

Next, ALP activity, an indicator of osteogenic differentiation [90,91], was evaluated. Consistent with previous studies demonstrating the osteogenic capabilities of CaP materials [92,93], our aim was to determine if these effects persist in the presence of fluoride and under varying topographical conditions. Fig. 9b depicts the ALP activity of SaOS-2 cells cultured on all tested CaP samples. The Flat control exhibited significantly elevated levels of ALP activity compared to the Ti control. Notably, most of the tested samples also showed higher ALP activity than the Ti sample, highlighting the potential to induce and support bone regeneration. While the effect of fluoride on osteoblast activity remains a subject of debate, literature reports suggests that fluoridated hydroxyapatite may slightly delay differentiation, an observation aligned with our findings for the 05F_AC or BM and the 10F_AC or BM samples [94,95]. Nevertheless, although these samples exhibited somewhat lower ALP activity at early time points, their values were comparable to those of the Ti control, indicating no adverse effects on osteogenic potential. This finding highlights the strong osteoinductive capacity of the CaP substrates, regardless of their nanotopographical variations. Notably, beyond in vitro analyses, prior investigations have demonstrated that nanostructured CaP materials, particularly those featuring needle-like nanoarchitectures, exhibit both osteogenic and osteoinductive properties under in vivo conditions [20,92]. Therefore, to fully explore these complex in vivo dynamics, future work is essential. Our next steps will include long-term biofilm formation assays and, ultimately, in vivo infection models to confirm the long-term clinical potential of these materials.

3.3.2. Cell morphology

Fig. 10 illustrates SaOS-2 cell adhesion and morphology after 4 h and 7 days of culturing. Additional time points are shown in Fig. S6.

After 4h of incubation, cell adhesion on the CaP substrates was lower than on the Ti surface, including the Flat sample. By day 7, both the Ti and Flat controls presented extensive surface coverage. Meanwhile, in agreement with the cell proliferation study, fewer cells were found in the nanostructured samples.

Across all evaluated time points, cells cultured on BM-treated samples (0F_BM, 05F_BM and 10F_BM) displayed a more spread morphology and well-defined cytoskeleton when compared to those cultured on AC discs (0F_AC, 05F_AC and 10F_AC). This difference was particularly pronounced for the 0F_AC sample, which exhibited the highest aspect ratio among all tested topographies and showed predominantly rounded cells at 4 h. These results align with the proliferation assay and indicate superior cytocompatibility for the low-temperature-treated samples.

3.3.3. Gene expression

To better understand the effect of the treated samples on SaOS-2 osteogenic differentiation, the expression of three key osteogenic genes (ALPL, RUNX2, and SPP1) was quantified by RT-qPCR at 1, 3, 7, and 14 days of culture. As shown in Fig. 11a, gene expression increased over time, with ALPL, RUNX2, and SPP1 upregulated at days 3, 7, and 14, respectively, and lower expression observed at day 1, consistent with the early-stage roles of RUNX2 and ALPL in osteoblast differentiation [96]. Both the substrate material and nanotopography influenced early differentiation, as evidenced by higher gene upregulation on 0F_AC and 0F_BM samples. ALP expression, an early osteogenic marker, peaked at day 7 across all substrates, in line with ALP activity assay results. Although minor differences related to topography were observed, all substrates markedly promoted early osteogenic differentiation, highlighting the central role of nanotopography alongside material properties.

Fig. 11 — (a) mRNA expression of osteogenic genes ALPL, RUNX2 and SPP1 of SaOS-2 cells cultured directly onto the CaP substrates for 1, 3, 7 and 14 days, determined by real-time PCR (n = 3). (b) Interaction of RAW 246.7 cells with treated discs up to 7 days in cell culture. i) mRNA expression of pro-inflammatory genes TNF, IL1B and IL6 of cells in the direct cell culture, determined by real-time PCR for 1, 3 and 7 days (n = 3). All values are relativized to values of cells at day 1. ii) Protein expression level after 7 days of direct cell culture on the treated discs, measured by inflammation antibody array. Values of protein signal are quantified by image analysis and relativized to control. In Fig. a) and bi), the same letter indicates no statistically significant differences for the same group at different time points while the same number denotes no statistically significant differences for each time point among all samples. (p < 0.05).

Parallel to these osteogenic effects, RT-qPCR analysis of relevant immunomodulatory genes of RAW 264.7 cells revealed the influence of nanostructure on inflammation (Fig. 11bi). As early as day 1, cells in contact with the nanostructured surfaces exhibited downregulation of IL-1β, whereas the Flat control, lacking nanotopography, displayed upregulation of all three cytokines, indicating a pro-inflammatory phenotype. The 0F_AC sample showed consistent downregulation of M1 markers across all timepoints, suggesting a pronounced anti-inflammatory effect, likely mediated by mechanotransduction through its nanostructures. Supporting this, previous studies on hydroxyapatite have reported elevated M2 marker expression after 3 days of incubation on nanoneedle structures [97]. In contrast, 10F_AC, 0F_BM, and 10F_BM samples, characterized by shorter needles and smaller inter-needle spacing, induced upregulation of TNF-α and IL-6, reflecting a transient pro-inflammatory phenotype that persisted until day 3 but declined significantly by day 7. This effect was particularly notable for IL-6, a later-phase cytokine with prolonged expression compared to TNF-α and IL-1β [98,99]. By day 7, all nanostructured surfaces promoted downregulation of inflammatory markers relative to the Flat control.

Overall, these results demonstrate that the nanostructured substrates not only enhance early osteogenic differentiation but also modulate macrophage inflammatory responses, highlighting the dual role of material topography in promoting bone regeneration while controlling inflammation.

3.3.4. Inflammatory antibody array

To evaluate the immunomodulatory effects of the different nanotopographies, the expression of inflammation-related proteins secreted by RAW264.7 cells at day 7 was analyzed using a mouse inflammatory antibody array (Fig. 11bii and Fig. S7). A total of 15 inflammatory proteins were quantified. Among them, IL-4, M-CSF, and MCP-1 are associated with the M2 reparative phenotype, whereas XCL1, CCL1, CX3CL1, CXCL6, IL-12p70, MIP-1γ, TNF RI, TNF RII, IFN-γ, IL-1β, and TNF-α correspond to the M1 pro-inflammatory phenotype. The results revealed a moderate upregulation of M2 markers, accompanied by a downregulation of several M1-associated proteins for all samples. Specifically, IL-4 and M-CSF were consistently upregulated across all conditions, including the Flat condition. This indicates enhanced macrophage polarization toward a regenerative profile, consistent with the immunomodulatory features of CDHA demonstrated in previous works [100,101]. In contrast, the decreased expression of TNF RI and TNF RII reflects a dampened inflammatory signal, consistent with the transcriptional repression of TNF-α.

Notably, the protein levels of IL-1β, IL-6, and TNF-α at day 7 correlated with the qRT-PCR data, both showing reduced expression relative to controls. This concordant decrease at the mRNA and protein levels strongly indicates that macrophages have transitioned from an acute inflammatory state toward a pro-healing, tissue repair phenotype.

3.3.5. In vitro co-culture evaluation

Coculture models represent a powerful tool for evaluating antimicrobial efficacy in a more physiologically relevant context. By incorporating both bacteria and osteoblastic cells, these systems more accurately mimic the clinical environment at the bone–implant interface, allowing simultaneous assessment of cellular responses and resistance to bacterial colonization [102]. Based on the previous results, 10F_AC and 10F_BM substrates were selected, together with the fluoride-free counterparts, based on their strong antibacterial performance and were co-cultured with SaOS-2 cells and P. aeruginosa using both pre- and post-implantation infection models. Flat and Ti discs were used as controls.

The CLSM images obtained from the pre-implantation infection model, in which SaOS-2 cells were cultured for 24 h on the substrates previously incubated with P. aeruginosa for 6 h, are presented in Fig. 12ai. Cell morphology was severely compromised under all conditions except for 10F_BM, which uniquely sustained well-spread cells despite bacterial challenge. In contrast, the remaining samples predominantly exhibited rounded cells, indicating cytoskeletal disruption. Cell adhesion was also significantly reduced on Ti and Flat controls, with a pronounced reduction in cell-covered area. These results are consistent with previous studies, which reported that the presence of P. aeruginosa can compromise SaOS-2 viability through the secretion of cytotoxic factors [103]. Cells without the incorporation of bacteria are shown in Fig. S8 for comparison.

Fig. 12 — Co-culture of *P. aeruginosa* and SaOS-2 cells on the nanostructured CaP discs, as well as Flat and Ti controls. a) Merged CLSM images of: i) a pre-implantation infection model, where the samples were first incubated for 6 h with *P. aeruginosa* and subsequently SaOS-2 cells were cultured for 24 h; or ii) post-implantation infection model, where SaOS-2 cells were first cultured for 24 h on the substrates, which were subsequently incubated for 6 h with *P. aeruginosa*. Actin filaments were stained with Alexa Fluor™ 546 phalloidin (orange fluorescence signal) and the nuclei with DAPI (blue fluorescence signal). (b) Orthogonal CLSM images showing simultaneous co-visualization of cells and bacteria stained with Alexa Fluor™ 546 phalloidin (orange fluorescence signal), the nuclei with DAPI (blue fluorescence signal) and SYTO-9 (green fluorescence signal). (c) Dead percentage of *P. aeruginosa* onto the different CaP substrates and the controls, for both the pre-implantation infection i) and the post-implantation ii) infection models. n = 3; ns and ∗ indicate significance at p > 0.05 and p ≤ 0.05, respectively.

CLSM images of the post-implantation infection model, in which SaOS-2 cells were first cultured for 24 h prior to the addition of P. aeruginosa for 6 h, are shown in Fig. 12aii. In this setup, previously adhered cells remained largely unaffected by bacterial exposure, with the exception of those on Ti and Flat surfaces, which exhibited severe morphological damage. Compared to the pre-implantation infection model, a greater number of cells were observed, displaying predominantly spread morphologies. Notably, treated substrates showed numerous well-adhered cells, indicating improved cytocompatibility. These findings confirm that Ti and Flat substrates are highly susceptible to bacterial-induced cytotoxicity under co-culture conditions, whereas nanotopographical surfaces, especially those incorporating fluoride, promote SaOS-2 adhesion and offer protection against bacterial colonization.

To gain insight into bacteria-cell interactions, bacterial localization analyses were performed, confirming that bacteria remained extracellular in both infection models (Fig. 12b). Finally, bacterial viability on the various substrates in both infection models was assessed using CLSM. As shown in Fig. 12c, nanostructured samples exhibited inhibitory effects on bacterial colonization across both models, with the fluoride-doped samples showing the strongest antibacterial activity. Notably, in the pre-implantation infection model (Fig. 12ci), differences between fluoride-doped and fluoride-free CaP samples were less obvious, with no statistically significant differences between them. However, both nanostructured CaP groups induced significantly greater bacterial death compared to the Flat and Ti control substrates.

In the post-implantation infection model (Fig. 12cii), bacterial death was also significantly higher on the nanostructured CaP substrates than on the controls. Moreover, fluoride-doped samples showed the highest antibacterial effect with statistically significant differences compared to undoped nanotopographies. Notably, a higher percentage of bacterial death was observed in both co-culture models compared to the bacterial monoculture system (section 3.2.1). This suggests that the bactericidal effect may be enhanced in the presence of cells, a phenomenon that warrants more in-depth investigation in future studies.

These co-culture results are of great interest as they directly address critical initial stages in implant-related infections. Specifically, the efficacy demonstrated in the pre-implantation infection co-culture setup directly relates to preventing intraoperative contamination, where bacteria are present immediately. Meanwhile, the superior performance observed in the post-implantation co-culture model offers particular relevance for mitigating infections arising from hematogenous spread, where bacteria arrive at an already colonized implant. Given that bacterial colonization of the biomaterial surface is recognized as the crucial first step towards the development of such infections, our results highlight a significant potential for preventing these common pathways of infection development.

In summary, this work provides a comprehensive cellular-level in vitro framework for understanding how nanotopographically engineered surfaces can favorably influence the “race for the surface” by promoting early host cell adhesion while simultaneously limiting initial bacterial colonization. Such nanostructured surfaces offer the dual benefits of modulating the host immune response to promote anti-inflammatory activity and support tissue integration [14] while also creating an environment where mammalian cells can progressively dominate the surface and inhibit bacterial proliferation [4]. However, although the co-culture model provides enhanced physiological relevance compared to conventional monoculture systems and enables mechanistic insight into bacteria–cell–material interactions, it does not fully capture the complexity of the in vivo environment. Translating these promising findings toward clinical application remains challenging, as in vivo settings introduce additional layers of complexity, such as immune responses, hemodynamic forces, dynamic protein adsorption, and long-term tissue remodeling processes, which cannot be fully captured in vitro. Moreover, the sustained antibacterial performance of such surfaces may be compromised by protein fouling or the accumulation of bacterial debris, which can mask nanotopographic cues and attenuate their functionality [104]. As a natural progression of this work, future research will include long-term biofilm assays and comprehensive in vivo infection models, which will be essential for validating the translational potential and long-term therapeutic efficacy of these biomaterial-based strategies.

4. Conclusions

In this study, we presented a biomimetic nanoengineering strategy for the fabrication of multifunctional CaP surfaces with enhanced antibacterial and osteogenic activities through controlled fluoride doping. By modulating the hydrolysis of α-TCP in fluoride-containing solutions, we were able to generate a series of nanostructured CDHA surfaces with tunable needle-like topographies and fluoride content. Such nanostructures were found to exhibit contact-dependent antibacterial activity, particularly against P. aeruginosa, while showing more limited efficacy against S. aureus. Fluoride incorporation clearly enhanced the antibacterial efficacy of the surfaces. This finding confirms a synergistic interplay between the mechanical action of the nanostructure and the chemical effect of fluoride.

Beyond antimicrobial activity, fluoride-modified surfaces supported cell adhesion, proliferation, and osteogenic differentiation, with particularly favorable responses for biomimetic samples. Moreover, the co-culture infection models, which simulate both pre- and post-implantation scenarios, confirmed the multifunctional nature of the fluoride-doped nanotopographies. They simultaneously promoted hist cell adhesion while inhibiting bacterial colonization through their enhanced bactericidal properties, a dual action that underscore their strong translational potential.

Overall, our approach demonstrates that the strategic integration of nanotopographical surface engineering and ion doping can yield CaP surfaces with dual antibacterial and osteogenic functionalities. This offers a promising pathway toward the development of next-generation bone graft biomaterials aimed at mitigating implant-associated infections while supporting bone regeneration in orthopedic and dental applications. Building on these findings, comprehensive in vivo studies will be essential to validate the translational relevance, long-term performance, and clinical potential of this approach.

CRediT authorship contribution statement

Carla Arca-Garcia: Writing – original draft, Investigation, Formal analysis, Data curation. Maria Godoy-Gallardo: Writing – original draft, Supervision, Methodology, Investigation, Data curation. Maria-Pau Ginebra: Writing – review & editing, Supervision, Methodology, Funding acquisition, Data curation, Conceptualization.

Ethics approval and consent to participate

The work does not involve research with human or animal subjects.

Declaration of competing interest

The authors declare the following financial interests/personal relationships which may be considered as potential competing interests:Maria-Pau Ginebra reports financial support was provided by European Research Council. Maria Godoy-Gallardo reports financial support was provided by Spanish Ministry of Science, Innovation and Universities. Maria-Pau Ginebra reports financial support was provided by Generalitat de Catalunya Ministry of Research and Universities. If there are other authors, they declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Acknowledgements

This work was funded by the European Union - ERC grant (BAMBBI; 101055053, doi: 10.3030/101055053). The research group is part of Maria de Maeztu Units of Excellence Programme CEX2023-001300-M/funded by MCIN/AEI/10.13039/501100011033, Spanish Ministry of Science and Innovation. The authors also acknowledge projects PID2022-142395OB-I00 and PID2023-148538OB-I00, funded by MICIU/AEI/10.13039/501100011033 and by FEDER, UE. M.-P.G. acknowledges the ICREA Academia award by the Generalitat de Catalunya. M.G.G. acknowledges the Spanish Ministry of Universities for the support received through the Ramon y Cajal Program (RYC2022-038428-I).

We would like to acknowledge BioRender for providing the tools to create some parts of the figures of this research article. Some images of the graphical abstract, Fig. 1, Fig. 2, Fig. 3, Fig. 5, Fig. 8, Fig. 9, Fig. 11, Fig. 12 were created in BioRender. Ginebra, M. (2025).

https://app.biorender.com/illustrations/68863daa0803117ec8fccea8?slideId=d22522ea-8fc5-4ccf-98f6-d71ba8731929.

Footnotes

Peer review under the responsibility of editorial board of Bioactive Materials.

^{Appendix A}

Supplementary data to this article can be found online at https://doi.org/10.1016/j.bioactmat.2025.12.026.

Contributor Information

Carla Arca-Garcia, Email: carla.arca@upc.edu.

Maria Godoy-Gallardo, Email: maria.godoy.gallardo@upc.edu.

Maria-Pau Ginebra, Email: maria.pau.ginebra@upc.edu.

Appendix A. Supplementary data

The following is the Supplementary data to this article:

Multimedia component 1

mmc1.docx^{(15.7MB, docx)}

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PERMALINK

Tailoring nanotopography and antibacterial properties of calcium phosphate bone grafts via fluoride incorporation

Carla Arca-Garcia

Maria Godoy-Gallardo

Maria-Pau Ginebra

Abstract

Graphical abstract

Highlights

1. Introduction

2. Materials and methods

2.1. Materials

2.2. Fabrication of nanostructured CDHA substrates

2.2.1. Powder preparation

2.2.2. Preparation of nanostructured CDHA discs

Fig. 1.

2.3. Physico-chemical characterization of the surfaces

2.3.1. Morphological analysis

2.3.2. Phase composition

2.3.3. Chemical characterization

2.3.4. Contact angle

2.3.5. Mechanical properties

2.4. Biological characterization of the nanostructured surfaces

2.4.1. Antimicrobial properties of the CDHA nanotopographies

2.4.1.1. Bacteria interaction with the CDHA nanotopographies

2.4.1.2. Bacterial morphology

2.4.2. Cell experiments

2.4.2.1. Cell proliferation and morphology

2.4.2.2. Alkaline phosphatase activity

2.4.2.3. Gene expression

2.4.2.4. Inflammatory antibody array

2.4.3. In vitro co-culture evaluation

2.4.3.1. Pre-implantation infection protocol

2.4.3.2. Post-implantation infection protocol

2.5. Statistical analysis

3. Results and discussion

3.1. Synthesis of F-doped nanotopographies and characterization

3.1.1. Morphological analysis

Fig. 2.

Fig. 3.

3.1.2. Phase characterization

Fig. 4.

3.1.3. Elemental composition

Fig. 5.

Fig. 6.

3.1.4. Contact angle analysis

Fig. 7.

3.1.5. Mechanical properties

3.2. Antibacterial activity

3.2.1. Early bacterial colonization

Fig. 8.

3.2.2. Bacteria morphology

3.3. Cell experiments

3.3.1. Cell proliferation and differentiation

Fig. 9.

3.3.2. Cell morphology

Fig. 10.

3.3.3. Gene expression

Fig. 11.

3.3.4. Inflammatory antibody array

3.3.5. In vitro co-culture evaluation

Fig. 12.

4. Conclusions

CRediT authorship contribution statement

Ethics approval and consent to participate

Declaration of competing interest

Acknowledgements

Footnotes

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Appendix A. Supplementary data

References

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