Tuning Anticancer Activity and Antimicrobial Response of ZnO Nanoparticles Through Halogenosilane Surface Modification

1. Introduction

The design of new inorganic nanomaterials with improved properties has started to take into account the surface modification through functionalization with organic capping agents. Such a strategy has become a central pursuit in nanomedicine, as it allows for combining therapeutic action with diagnostic or imaging capabilities [1]. Within this context, zinc oxide nanoparticles (ZnO NPs) have emerged as a rare case in which a single material spans many field of research and scientific disciplines. Features like wide bandgap, strong exciton binding energy and relatively simple preparation methods make ZnO NPs attractive for applications ranging from electronics to catalysis, but also for biomedical uses [2,3,4,5,6,7].

Pristine ZnO NPs tend to agglomerate in aqueous media and may release Zn²⁺ ions uncontrollably, raising concerns over reproducibility and safety [8]. Consequently, surface modification has started to be adopted as a useful strategy to improve colloidal stability and to modulate biological interactions. In the realm of surface modification, the approach of organosilane functionalization stands out, because silanes can form robust covalent bonds with hydroxylated ZnO surfaces, imparting stability and allowing fine-tuning of surface polarity [9,10]. In this way, silanes are not merely stabilizers, but act as chemical tools to reshape ZnO’s physico-chemical and biological behavior.

The biological performance of ZnO is strongly influenced by particle size and morphology. Smaller ZnO NPs generally exhibit higher surface reactivity, enhanced cellular uptake and stronger antimicrobial and anticancer activities, though sometimes at the cost of cytotoxicity to healthy cells [11,12,13,14]. For instance, hepatocyte and mesenchymal stem cell studies have revealed clear size-dependent changes in oxidative stress and gene expression of genes related to senescence [15,16]. Morphological variations, such as nanosheets compared with flower-like structures, can also influence biological activity. For instance, sheet-like ZnO was shown to maintain dispersibility and superior antibacterial efficacy even under varying conditions [17].

The importance of defects has also been emphasized. Oxygen vacancies are known to facilitate ROS generation, sometimes even in the absence of light [18]. Intentional doping with gallium [19], magnesium [20] or silver [21] has been exploited to increase defect concentrations, and thus, amplify cytotoxic and antibacterial effects. The ZnO’s performance consistently highlight that size, shape and defect chemistry are intertwined with surface chemistry [22,23].

Organosilanes have become a central subject in tailoring ZnO for biomedical applications. For example, (3-glycidyloxypropyl)trimethoxysilane (GPTMS) coatings were shown to enhance colloidal stability, while maintaining antibacterial activity against E. coli and S. aureus [24]. Increasing GPTMS content even led to smaller particle sizes, with enhanced anticancer effects against ovarian cancer cells through ROS-mediated apoptosis and autophagy [24]. Influence of size and surface modification of GPTMS-modified ZnO NPs has been highlighted by increased antibacterial activity [25]. GPTMS-modified ZnO also retained antibacterial activity against multidrug-resistant Enterobacteriaceae, with improved stability during storage [26]. ZnO quantum dots capped with aminopropyltriethoxysilane (APTES) have shown promise in biosensing, bioimaging and antimicrobial activity, owing to their enhanced fluorescence and stability [27,28].

These examples are thus coming to validate the paradigm controlled surface–controlled performance, showing how organosilane functionalization may be a powerful lever for tuning ZnO NPs [29]. Yet, one avenue has remained largely unexplored, that is, the use of halogenosilanes. Functional groups such as chloropropyl-, bromopropyl- and iodopropyl attached to the trimethoxysilane moiety could, in principle, introduce distinctive surface characteristics. The presence of halogen atoms may influence the polarity, hydrophobicity and defect states, which could alter colloidal stability, ROS generation and interactions with cancer or microbial cell membranes. Halogenated silanes have already been employed in other material systems, often conferring higher reactivity or antimicrobial potential [30,31]. In biomedical surfaces, silane chemistry has also been extended to implantable biomaterials. Somasundaram reviewed how silanes, often in combination with metallic biomaterials, can improve bioactivity and antimicrobial resistance in implant coatings [32]. More recently, Morselli and co-workers demonstrated that sprayable PEG-silane/ZnO nanocomposite coatings can successfully prevented microbial colonization of titanium implants [33]. Complementary strategies further illustrate the breadth of ZnO nanomaterials. Halogenosilane green synthesis approaches have produced ZnO NPs with antibacterial activity and acceptable cytocompatibility [34]. Clinical relevance has also been underscored by in vitro studies, in which ZnO NPs displayed both antibacterial and cytotoxic properties, emphasizing the importance of precise surface engineering to maximize benefit while minimizing risk [35].

However, to the best of our knowledge, no systematic study has addressed surface modification of ZnO NPs with halogenosilanes for anticancer and antimicrobial applications. This knowledge gap motivates the present work and herein we report on the first introduction of halogenosilane species, such as 3-chloropropyltrimethoxysilane (CPTMS), 3-bromopropyltrimethoxysilane (BPTMS) and 3-iodopropyltrimethoxysilane (IPTMS) into the surface modification practice of semiconductor ZnO NPs.

To expand the diversity of ZnO-based therapeutics we explore the ability of a new family of halogenosilane-modified ZnO nanoparticles to act as anticancer and antimicrobial agents, aiming to evaluate their therapeutic effectiveness. In this work, particular emphasis is placed on their syntheses, characterization, and biological evaluation as anticancer and antimicrobial agents.

2. Results and Discussion

2.1. Synthesis and Morpho-Structural Characterization

The preparation of the new series of ZnO NPs modified with the halogenosilane species (3-chloropropyl)trimethoxysilane (CPTMS), (3-bromopropyl)trimethoxysilane (BPTMS) or (3-iodopropyl)trimethoxysilane (IPTMS) was performed by the in situ chemical precipitation method, starting from zinc(II) acetate dihydrate, fixed amounts of halogenosilanes corresponding to 10% Si/Zn molar ratio, and potassium hydroxide, in ethanol under reflux for 3 h. The choice of using the 10% Si/Zn molar ratio was dictated by our previous observation that this was the maximum limit for reaching the highest surface coverage, and hence, the smallest ZnO NPs, upon surface modification with other organosilanes, such as (3-glycidyloxypropyl)trimethoxysilane (GPTMS) [24], (3-trimethoxysilyl)propylmethacrylate (MPS) [36], or vinyltrimethoxysilane (VTMS) [37,38]. Besides this, when it comes about their application in the biomedical field, the anticancer and antimicrobial properties were found to be size-dependent and, for instance, the smallest ZnO NPs, obtained by GPTMS-modification at the 10% Si/Zn ratio, displayed the best antibacterial performances against E. coli and S. aureus [24].

According to the proposed mechanism (Scheme 1), the formation of halogenosilane-modified ZnO NPs involves hydrolysis, in the basic ambient assured by potassium hydroxide, of zinc(II) acetate to form zinc(II) hydroxide (step 1), concomitantly with the hydrolysis of halogenosilane molecules to generate the halogensosilanol species (step 2) and, finally, the self-condensation reaction between the halogenosilanol species and zinc(II) hydroxide, in order to promote the formation of ZnO particles coated with a silica network bearing the pendant halogenopropyl groups (step 3).

X-ray diffraction (XRD) analysis allowed to confirm the presence of zincite phase of ZnO in both unmodified and halogenosilane-modified samples, crystallizing in the hexagonal P63mc space group. The XRD patterns of all synthesized ZnO samples reveal the presence of the characteristic crystallographic planes (100), (002), (101), (102), (110), (103), (200), (112), (201), (004) and (202) in the 30‒80° 2θ range (Figure 1). In the case of unmodified ZnO NPs, the XRD pattern shows well resolved and more intense diffraction peaks. Upon ZnO surface coating with halogenosilanes, the diffraction peaks undergo a significant intensity decrease and broadening, as a consequence of crystallinity reduction, along with particles downsizing. In fact, as noted in Table 1, the crystallinity reduction occurs from 59.16% in unmodified ZnO NPs to 49.75% in iodosilane-modified ZnO NPs, whereas the average crystallite size decreases from ca. 21 nm in unmodified ZnO NPs to around 7 nm in all halogenosilane-modified ZnO NPs, thus highlighting the important size reduction effect of surface modification of ZnO with these types of organosilane surfactants. It is also worthy of mention that the observed differences in the crystallite size could also be due to eventual crystal microstructure changes induced by several factors, such as inhomogeneous strain, point defect, bending, dislocation generation and diffusion [39,40].

Transmission electron microscopy (TEM) allowed to reveal the clear influence of halogenosilane surface modification on the morphology and size of the synthesized ZnO NPs. The unmodified ZnO NPs consisted of irregularly faceted and nearly spherical particles (shape factor close to 1), which form dense agglomerates, with an average particle size of 30.77 ± 0.35 nm (Figure 2, sample ZnO), which is indicative of an extensive coalescence due to high surface energy and the absence of stabilizing agents [41,42]. After modification with chloropropylsilane, the mean particle size decreased markedly to 14.62 ± 0.28 nm, and the particles appeared more discrete and uniformly dispersed, while their shape started to undergo a slight distortion from the spherical one (shape factor close to 0.9) (Figure 2, sample ZnO_c). The reduction in particle size and narrowing of the distribution suggest that the chloropropyl moieties strongly interact with surface hydroxyl groups, effectively suppressing secondary growth during nucleation and crystallization, as the silane species may directly participate in the condensation reaction with the zinc hydroxide and zinc oxide species. This leads to a subsequent limiting effect of the self-condensation of zinc hydroxide in the primary growth of ZnO crystallites and in their agglomeration [43,44]. The bromopropylsilane-modified ZnO NPs exhibited the smallest average size (8.31 ± 0.15 nm), with nearly elongated shapes and well-dispersed nanoparticles (Figure 2, sample ZnO_b). This indicates that bromopropysilane provided the most efficient capping among the three halogenosilane derivatives, combining adequate surface reactivity with enhanced steric hindrance. In contrast, the iodopropylsilane-modified ZnO NPs displayed a slightly larger mean diameter (11.62 ± 0.42 nm) and a broader distribution (Figure 2, sample ZnO_i), implying that the bulky iodine substituent reduced surface-binding strength and capping efficiency.

Overall, halogenosilane modification significantly improved nanoparticle dispersion and reduced agglomeration, with the effectiveness of these modifiers following the trend ZnO_b > ZnO_c > ZnO_i > ZnO. The observed reduction in particle size and enhancement in morphological uniformity are consistent with earlier studies, which demonstrate that silane coupling agents can effectively passivate ZnO surfaces, disrupt agglomeration and limit the particle growth during chemical precipitation synthesis [45,46,47]. This finding supports the interpretation that the interplay between surface-bonding strength (Si–O–Zn formation) and steric effects of the halogen substituent governs growth inhibition efficiency. The smaller and more reactive halogens (Cl and Br) favor stronger surface anchoring, while bulkier iodine reduces capping uniformity, leading to partial coalescence. The observed differences in size and morphology may have a substantial influence on the physicochemical characteristics of the modified ZnO NPs, as well as their potential biomedical applications.

Quantitative EDX compositional analysis was conducted to further support successful halogenosilane modification of ZnO NPs (Figure 3). While the unmodified ZnO sample exhibits only Zn and O signals, upon modification all functionalized samples show additional peaks corresponding to Si and the respective halogen (Cl, Br, I). The detection of Si in all modified samples verifies the presence of silane coupling on ZnO surfaces, while the simultaneous detection of halogens supports retention of the Si–propyl‒X functionality post-modification. The decreasing Zn atomic fraction relative to the control reflects surface coverage by lighter elements (Si, C, halogen) and corroborates the formation of an organic–inorganic hybrid shell. Moreover, the relative atomic percentages suggest variable grafting efficiencies, governed by both the reactivity of the Si–O bond and the steric size of the halogen substituent.

Fourier transform infrared spectroscopy (FTIR) was employed to verify the success of halogenosilanes surface coating of ZnO NPs. The FTIR spectra of free halogenosilanes, as well as those of unmodified and halogenosilane-modified ZnO NPs are given in Figure 4. In the FTIR spectra of free halogenosilanes, the strong absorption bands located at 1074-1076 cm^-1 and 800-804 cm^-1 were assigned to the stretching vibration of C‒O bonds from methoxy groups and the symmetric stretching of Si‒O‒C bonds, respectively. In all halogenosilane-modified ZnO NPs, these bands completely disappear, as a consequence of the hydrolysis of methoxy groups bound to silicon. The new strong band that appeared at 880-881 cm^-1 in all halogenosilane-modified ZnO NPs indicates that Zn‒O‒Si bonds, as well as Si‒O‒Si bridges, are formed through the self-condensation reaction of the halogenosilanol species surrounding the ZnO particles [48], thus giving them a core-shell structural model with the halogens oriented outwardly. The strong bands clearly observed at 756-769 cm^-1 for the C‒halogen bonds in the free halogenosilanes, are no longer evident in the halogenosilane-coated ZnO NPs, and are likely overlapped with the strong bands already detected for the formation of Zn‒O‒Si bonds and Si‒O‒Si bridges.

In the FTIR spectra of both unmodified and halogenosilane-modified ZnO NPs, the broad bands located in the range 3376-3384 cm^-1 were assigned to the stretching vibration of hydroxyl groups of physisorbed water molecules. In the case of unmodified ZnO NPs, the weak broad band found at 892 cm^-1 was assigned to the bending mode of carbon dioxide molecules, possibly physisorbed from air on the zinc (II) ions from the ZnO surface [49,50]. Also, in all ZnO samples two strong bands observed at 1569 and 1416 cm^-1 in unmodified ZnO NPs, 1559 and 1418 cm^-1 in CPTMS-modified ZnO NPs, 1570 and 1418 cm^-1 in BPTMS-modified ZnO NPs, and 1559 and 1418 cm^-1 in IPTMS-modified ZnO NPs, were assigned to the asymmetric and symmetric stretching of remaining acetate groups from the initial zinc(II) precursor, which, after repeated washings, could not be removed, thus giving rise to consider that the carboxylate groups were covalently attached onto the zinc(II) ions [51].

2.2. Biological Properties

2.2.1. Cytotoxic Activity

ZnO NPs have received much attention for their applications in cancer therapy. However, the underlying molecular mechanisms behind their anticancer activity remain unclear. It has been reported that ZnO NPs induce apoptosis mediated by reactive oxygen species generation [52]. The cytotoxicity of the halogenosilane derivatives have been reported to depend mostly on their physicochemical properties such as shape, size, and surface modifications. Size is in fact a very important factor, influencing the uptake and cytotoxicity [53]. Results of the cytotoxic effect of the unmodified and halogenosilane-modified ZnO NPs on the prostate cancer cells (PC3 and 22Rv1), normal prostate cells (RWPE) and fibroblasts (HDF) using the MTT colorimetric assay are presented in Table 2. After 24 h incubation with the cells, the halogenosilane-modified ZnO NPs showed a strong cytotoxic effect on both cancer cell types and the unmodified ZnO showed lower cytotoxicity. In the normal prostate cells and fibroblasts, all ZnO NPs samples were slightly less active, in particular for the unmodified ZnO and ZnO_b which indicate some selectivity for cancer cells.

2.2.2. Cellular Uptake by ICP-MS

The cellular uptake can be influenced by the size and the surface modification of the ZnO NPs. As can be observed in Figure 5, the cellular uptake in the PC3 cells (in terms of Zn levels) of unmodified ZnO was low compared to the halogenosilane-modified ZnO NPs in particular ZnO_b and ZnO_c.

2.2.3. Oxidative Stress by ROS

ZnO NPs have the unique ability to induce oxidative stress in cancer cells, which has been found to be one of the mechanisms of cytotoxicity [54]. Reactive oxygen species (ROS) generation and membrane lipid peroxidation was identified as a likely mechanism of ZnO NPs toxicity [24]. The production of ROS (mainly peroxides) was measured using the cell-permeant indicator H₂DCF-DA. The fluorescence signal of DCF was measured after 3 and 24 h. As can be observed from Figure 6, production of ROS is time and concentration dependent. After 24 h, an important increase of DCF fluorescence was observed in particular for ZnO_b.

2.2.4. Mechanisms of Cell Death

2.2.4.1. Apoptosis (Caspase 9 Activity)

The oxidative stress could lead to cell membranes damage and, depending on that, apoptosis may be triggered which leads to cell death. Apoptosis is believed to be, for some nanoplatforms, the major mechanism of cell death in the cytotoxic response of ZnO NPs by the activation of caspases [55]. In this work, apoptosis was identified by measuring the level of caspase 9, the central initiator caspase.

Results in Figure 7 show that all the ZnO NPs induce apoptosis in PC3 cells, in a dose dependent way, with emphasis on ZnO. The cytotoxicity results indicate that besides apoptosis, additional mechanisms of cell death are triggered since higher levels of apoptosis was observed for ZnO, which shows the lower level of cellular cytotoxicity (Table 2). On the other hand, ZnO_b shows the higher level of cellular cytotoxicity and lower levels of apoptosis. Also, the apoptosis levels of ZnO NPs do not correlate with the cellular uptake as can be observed from Figure 5 that show higher uptake for ZnO_b and ZnO_c. In the case of ZnO_b, ROS production seemed to mediate cell death.

These observations suggest that different mechanisms may contribute to cell death depending on nanoparticle surface properties, with unmodified ZnO favouring apoptosis-related pathways, while halogenosilane-modified systems, particularly ZnO_b, may involve additional ROS-driven or non-apoptotic mechanisms.

2.2.4.2. Morphological Features by SEM

The morphological features of cells upon exposure to ZnO NPs were examined through the analysis of scanning electron microscopy (SEM). Cell-surface changes, including membrane blebbing and loss of cellular features were observed (Fig. 8). Figure 8 shows untreated cells (control (CTR), Fig. 8a) presenting normal cellular morphology, including migrating cells (m) and dividing cells (d). In Fig. 8b (ZnO), cells appear rounded (r) and are surrounded by membrane blebs (b), features compatible with apoptotic phenomena. In Fig. 8c (ZnO_b), cells extend membrane projections (p), although often without the clear polarity observed in migrating control cells. Fig. 8d (ZnO_c) shows membrane activity similar to that observed in Fig. 8c. These observations suggest a marked contribution of apoptosis in ZnO-treated cells. In contrast, ZnO_b- and ZnO_c-treated cells still show signs of cellular activity, but these appear substantially altered and are probably dysfunctional when compared with CTR cells. Results are in agreement with those of caspase 9 activity assays (Fig. 7) that show the ability of ZnO to activate caspase.

These morphological alterations are consistent with the biochemical assays and further support the involvement of oxidative stress and apoptosis-related processes. Moreover, the differences observed between ZnO and halogenosilane-modified samples reinforce the role of surface functionalization in modulating nanoparticle–cell interactions.

2.2.5. Antimicrobial Activity of ZnO and Halogenosilane-Modified ZnO Nanoparticles

The antimicrobial activities of ZnO and halogenosilane-modified ZnO NPs was assessed towards the Gram-negative Pseudomonas aeruginosa, Escherichia coli, and Burkholderia contaminans, the Gram-positive Staphylococcus aureus, and the fungi Candida albicans and C. glabrata. Attempts to determine the MIC values based on the measurement of absorbance of microbial suspensions were hindered by the insoluble nature of the ZnO NPs. Therefore, we opted to incubate the microbial cultures in the presence of the ZnO and halogenosilane-modified ZnO NPs at final concentrations of 1000, 1250 and 1500 µg/mL for 24 hours and enumerate the CFUs formed after 24 h of spreading 100 µL cultures on the surface of LB (bacteria) or PDA (fungi). The minimal concentrations of ZnO and halogenosilane-modified ZnO NPs, for which no microbial growth was registered, are shown in Table 3.

Concentrations up to 1500 µg/mL of the the ZnO and halogenosilane-modified ZnO NPs were not bactericidal for P. aeruginosa and B. contaminans, and were also not fungicidal to C. albicans and C. glabrata (Table 3). For E. coli the halogenosilane-modified ZnO NPs were more effective than the ZnO NPs. This was also the case for S. aureus, with the exception of ZnO-b, which presented a minimal bactericidal concentration identical to that of ZnO NPs.

ZnO nanoparticles antimicrobial activities have been tested by several authors. For instance, ZnO NPs MIC values of 62.5 µg/mL towards the Gram-negative E. coli and Salmonella typimurium were reported by Zhong et al. [56]. These authors reported MIC values of 31.25 µg/mL for the Gram-positive B. subtilis and E. faecalis. Interestingly, these authors also determined the percentage of viability of the same cultures and values of 5-10% viability were reported for ZnO NPs concentrations of 500 µg/mL. In our work, bactericidal activity (0 % viability) was only achieved for concentrations of at least 1250 µg/mL (Table 3). Zhong et al. have not reported minimal bactericidal concentrations for the bacterial species tested.

Several mechanisms underlying the antimicrobial activity of ZnO nanoparticles have been described, including the generation of oxygen reactive species (ROS) in the presence of light. Additionally, nanoparticles can be adsorbed by the bacterial membrane, compromising its integrity and permeability, facilitating the internalization of nanoparticles that can lead to elevated and toxic Zn²⁺ concentrations within the bacterial cytoplasm [57]. In the present work, the mechanisms underlying the microbiocidal activity of the ZnO and halogenosilane-modified ZnO NPs was not addressed.

These results suggest that halogenosilane surface functionalization may influence the antimicrobial behavior of ZnO nanoparticles, particularly against E. coli and S. aureus, possibly through combined effects of particle size reduction and surface chemistry modification. This behavior, together with the enhanced anticancer activity, supports the idea that biological responses to ZnO nanomaterials can be modulated through rational surface engineering.

3. Materials and Methods

3.1. Reagents

Zinc acetate dihydrate (purity > 98%), potassium hydroxide (purity ≥ 85%), ethanol (purity = 98%), (3-chloropropyl)trimethoxysilane (purity ≥ 97%), (3-bromopropyl)trimethoxysilane (purity ≥ 97%) and (3-iodopropyl)trimethoxysilane (purity ≥ 97%) were purchased from Merck (Darmstadt, Germany) and used as received.

3.2. Synthesis of Halogenosilane-Modified ZnO NPs

In a 50-mL round-bottom flask, 10 mL of ethanol was introduced and heated to a temperature of 45–50 °C, under constant stirring, using a heating plate with magnetic stirring. This step ensures a homogeneous reaction medium and favors the interaction between ethanol and the chemical precursors. The halogenosilane species (3-chloropropyl)trimethoxysilane (CPTMS), (3-bromopropyl)trimethoxysilane (BPTMS) or (3-iodopropyl)trimethoxysilane (IPTMS) was added to the ethanol solution, in a 10% Si/Zn molar ratio. After the addition of halogenosilane species to ethanol, the temperature of the solution was progressively increased to 60 °C and maintained constant for 10 minutes, to allow partial hydrolysis of the halogenosilane species and its activation for the subsequent reaction with zinc acetate. 2.45 g of zinc acetate dihydrate were added to the halogenosilane-containing ethanol solution, under continuous magnetic stirring. Zinc acetate serves as a precursor for the formation of ZnO nanoparticles, and its interaction with silanes modifies the surface properties of the obtained particles. Separately, 1.25 g of potassium hydroxide (KOH) were dissolved in 5 mL of ethanol, under constant stirring at room temperature. This solution was subsequently used to induce the precipitation of ZnO and the formation of ZnO NPs. The KOH solution was added dropwise over the zinc acetate and halogenosilane solution, under constant stirring. This process is necessary to control the nucleation and growth of nanoparticles, avoiding the formation of ZnO aggregates. After completing the KOH addition, the solution was maintained under magnetic stirring and reflux for 3 hours, in order to ensure the formation and stabilization of ZnO NPs, as well as to functionalize their surface with the hydrolyzed halogenosilane derivatives. The obtained halogenosilane-modified ZnO NPs were collected by centrifugation (9000 rpm, 5 min, 10 °C) and washed successively with three portions (5 mL) of ethanol. The final products were dried in an oven at 100 °C for 2 hours. While the unmodified ZnO NPs sample is hereafter denoted as ZnO, while the ZnO NPs samples modified with CPTMS, BPTMS and IPTMS are denoted as ZnO_c, ZnO_b and ZnO_i, respectively.

3.3. Nanoparticles Characterization

3.3.1. XRD Analysis

X-ray diffraction (XRD) patterns were recorded on ground powders with an Empyrean (PANalytical, the Netherlands) diffractometer, using a Cu K_α1 (λ = 1.540598 Å) radiation, equipped with 2xGe (220) hybrid monochromator for Cu and PIXcel3D detector. The analyses were performed using Bragg-Brentano geometry (“theta-2theta”) for angles between 10 and 80° 2θ degrees, with a scan step increment of 0.04° and acquisition time on step size of 3 s. Sample preparation: a small amount of each sample was mounted on a single crystal silicon holder, which does not give diffraction interferences in the analysis range for the radiation emitted by a Cu anode. The sample was gently pressed on the holder so that the analyzed area be flat. For the identification of cell parameters and mean crystallite size, Rietveld refinement was performed in PANalytical High Score Plus v3.0e software. The agreement indices considered for the refinement are R_expected, R_profile, Weighted R profile and Goodness of fit. In Table 1, relevant crystallographic data for unmodified and halogenosilane-modified ZnO NPs are provided.

Table 1. Lattice parameters obtained from XRD analyses of the obtained ZnO NPs.

Crystallographic data	ZnO	ZnO_c	ZnO_b	ZnO_i
Crystallinity Degree (%)	59.16	53.10	52.86	49.75
ICDD PDF4+	04-007-1614	04-006-9717	04-007-1614	04-006-9717
Phase	Zincite	Zincite	Zincite	Zincite
Crystallization group	P63mc	P63mc	P63mc	P63mc
Compositional percentage (%)	100	100	100	100
a [Å]	3.25244	3.25418	3.254538	3.253683
b [Å]	3.25244	3.25418	3.254538	3.253683
c [Å]	5.21007	5.21225	5.212538	5.211145
alpha [°]	90	90	90	90
beta [°]	90	90	90	90
gamma [°]	120	120	120	120
Volume [Å^3]	47.73029	47.80137	47.81439	47.77649
Crystallite size (A)	20.970	7.388	6.919	6.998
Crystal System	Hexagonal	Hexagonal	Hexagonal	Hexagonal
R expected	6.3157	6.14177	6.11882	6.26389
R profile	4.75272	5.06297	5.21976	5.10729
Weighted R profile	6.30025	6.40155	6.64908	6.660314
Goodness of fit	0.99511	1.08638	1.18083	1.13154

Table 1. Lattice parameters obtained from XRD analyses of the obtained ZnO NPs.

Crystallographic data	ZnO	ZnO_c	ZnO_b	ZnO_i
Crystallinity Degree (%)	59.16	53.10	52.86	49.75
ICDD PDF4+	04-007-1614	04-006-9717	04-007-1614	04-006-9717
Phase	Zincite	Zincite	Zincite	Zincite
Crystallization group	P63mc	P63mc	P63mc	P63mc
Compositional percentage (%)	100	100	100	100
a [Å]	3.25244	3.25418	3.254538	3.253683
b [Å]	3.25244	3.25418	3.254538	3.253683
c [Å]	5.21007	5.21225	5.212538	5.211145
alpha [°]	90	90	90	90
beta [°]	90	90	90	90
gamma [°]	120	120	120	120
Volume [Å^3]	47.73029	47.80137	47.81439	47.77649
Crystallite size (A)	20.970	7.388	6.919	6.998
Crystal System	Hexagonal	Hexagonal	Hexagonal	Hexagonal
R expected	6.3157	6.14177	6.11882	6.26389
R profile	4.75272	5.06297	5.21976	5.10729
Weighted R profile	6.30025	6.40155	6.64908	6.660314
Goodness of fit	0.99511	1.08638	1.18083	1.13154

The average crystallite size (L) of synthesized ZnO NPs was estimated by applying Scherrer’s equation (1):

$$\mathit{B}\left(2\theta \right)={\displaystyle \frac{\mathit{K}\cdot \mathit{\lambda }}{\mathit{L}\cdot cos\theta }}$$

(1)

where B is the full width at half maximum intensity (FWHM), λ is the X-ray wavelength, θ is the diffraction angle and K is the Scherrer’s constant, whose value for spherical particles is 0.89 [58].

3.3.2. TEM Analysis

Transmission electron microscopy (TEM) was conducted to analyze the morphology and size distribution of both unmodified and halogenosilane-modified ZnO NPs. The samples were examined using a JEOL JEM-1400 transmission electron microscope (JEOL, Japan) operated at 80 kV accelerating voltage, equipped with a Quemesa CCD camera (Olympus Soft Imaging Solutions). For statistical analysis, at least 100 nanoparticles were measured per sample. Using the ImageJ software (https://imagej.nih.gov/ij/), the crystal outlines were digitized and their dimensions measured. The resulting data were afterwards used to generate histograms in Origin software, illustrating the average size of the nanoparticles. For each sample, a 1 mg/mL dilution in absolute ethanol was prepared and thoroughly dispersed using a Sonorex Digital 10 P ultrasonic bath (Bandelin electronic, Heinrichstraße, Berlin). Subsequently, working in duplicate, a droplet of the nanoparticle suspension was applied onto 200 mesh formvar-coated grids and allowed to air-dry.

3.3.3. SEM-EDX Analysis

The morphology and chemical composition of ZnO samples were analyzed through scanning electron microscopy (SEM) using the FEI Quanta 200 system (FEI Company, Hillsboro, OR, USA) operating at 15 kV, coupled with an energy dispersive X-ray spectroscopy (EDX) system. Specimens were prepared by dispersing the samples by sonication in ethanol and by depositing a few drops of the suspensions on carbon-coated grids. To minimize charging effects during SEM examination, the samples were coated with a thin gold layer deposited by sputtering using an SPI-Module system (SPI Module™ Supplies, West Chester, PA, USA).

3.3.4. FTIR Analysis

Fourier transform infrared spectra (FTIR) were collected using a Nicolet iS50 FTIR spectrometer (Thermo Scientific, Waltham, MA, USA) equipped with a built-in ATR accessory, DTGS detector and KBr beam splitter. A total of 32 scans were added in the range 4000–400 cm⁻¹, with a resolution of 4 cm⁻¹.

3.4. Biological Assays

3.4.1. Anticancer Activity

PC3 (PSA-, AR-) (ATCC, CRL-1435) and 22Rv1 (PSA+, AR+) (ATCC, CRL-2505) human prostate cancer cells were cultured in RPMI medium supplemented with 10% fetal bovine serum (FBS) (Gibco, Thermo Fisher Scientific, USA), RWPE-1 normal human prostate cells (ATCC, CRL-3607) were cultured in keratinocyte serum-free medium (K-SFM) supplemented with bovine pituitary extract (BPE) (0.05 mg/ml), epidermal growth factor (EGF) (5 ng/ml) and 10% FBS, and the human dermal fibroblasts (HDF) (Sigma-Aldrich) were cultured in fibroblast growth medium (Sigma-Aldrich). Cultures were maintained in an incubator (Heraeus, Germany) with humidified atmosphere at 37 °C and 5% CO₂. The cytotoxic activity of the halogenosilanes-modified ZNO NPs was evaluated by the MTT colorimetric assay as previously described [24]. For the assays, cells (1‒2x10⁴ cells/200µL) were suspended in culture medium, seeded into 96-well plates and incubated for 24 h. A stock solution for each nanoplatform was prepared in PBS at a concentration of 1 mg/mL and sonicated for 5 min. Then, working solutions of 1, 5, 10, 20, 50, 100 and 200 μg/mL were prepared in complete media and added to the cells. Untreated cells were also included as the negative controls. After treatment with the NPs, the cell culture medium was replaced by 200 μL of MTT solution in PBS (0.5 mg/mL) followed by 3 h incubation at 37 °C. The resulting formazan product was solubilized with 200 μL DMSO and the absorbance measured at 575 nm. Each experiment was repeated at least twice, and each concentration was tested with at least six replicates. The IC₅₀ values were calculated from dose-response curves using the GraphPad Prism software (vs. 6.0).

3.4.2. Analysis of Cellular Zinc Content by ICP-MS

PC3 cells (~10⁶ cells) were seeded into T25 cell culture flasks with RPMI medium and in-cubated for 24 h. After, the ZnO-NPs were added to the cells at their IC₅₀ (µg/ml) in medium and incubated for 24 h at 37 °C. After treatment, the cell pellets obtained by centrifugation, were digested in 100 µL of HNO₃ combining ultrasound (60 min at 60 °C) and microwave (350 W, 15 s) radiation for Zn quantitation by a Thermo X-Series Quadrupole ICP-MS Thermo Scientific, Rockford, IL, USA as previously described [59].

3.4.2. ROS Production

The production of ROS (mainly peroxides) was measured using the fluorescent probe H₂DCF-DA (2',7'-dichlorodihydrofluorescein diacetate) as previously described [60]. Briefly, PC3 cells (~ 2 × 10⁴/200 µL) in RPMI medium with 10% FBS (Gibco) were seeded in 96-well plates and left to adhere overnight. Then, the medium was replaced with the ZnO-NPs suspensions in complete RPMI medium at 5, 10, 20, 50 and 100 µg/mL and left for 24 h incubation. After, the medium was replaced by a solution of 10 μM H₂DCF-DA in colorless FluoroBrite™ DMEM (Gibco) and cells were incubated for 30 min. at 37 °C. The probe solution was removed and substituted by the colorless medium. Dichlorofluorescein (DCF) fluorescence was measured at 492 nm excitation and 517 nm emission along the time up to 24 h using a Varioskan LUX scanning multimode reader (ThermoFisher Scientific). Results (mean ± SD) are expressed as arbitrary fluorescence units. The intensity of the resulting fluorescence is directly proportional to the amount of ROS present in the cells.

3.4.3. Apoptosis Assay

The mechanism of cell death by apoptosis was evaluated by a Caspase-Glo® 9 luminescent assay following the protocol of the supplier (Promega, Madison, WI, USA). The assay provides a luminogenic caspase-9 substrate in a buffer system optimized for caspase activity, luciferase activity and cell lysis. The luminescent signal produced by the luciferase reaction was measured using a Varioskan LUX scanning multimode reader (ThermoFisher Scientific). The signal generated is proportional to the amount of caspase activity present.

3.4.4. SEM Studies with Cells

PC3 cells were seeded over glass lamellae in 6-well plates at a density of 2 × 10⁵ cells in 2 mL cell medium and allowed to adhere for 24 h. After, the ZnOs were added to the cells in medium at a concentration corresponding to their IC₅₀ values, 66, 20 and 34 μM, respectively for ZnO, ZnO_b and ZnO_c .After 24 h incubation, the medium was discarded and the cell-covered glass lamellae were treated following a previously described method [61]. The samples were observed and photographed using a JEOL 5400 scanning electron microscope.

3.4.5. Antimicrobial Activities of ZnO and Halogenosilane ZnO NPs

The antimicrobial activity of the ZnO, ZnO_c, ZnO_b and ZnO_i nanoparticles was assessed towards the Gram-negative Pseudomonas aeruginosa, Escherichia coli and Burkholderia contaminans, the Gram-positive Staphylococcus aureus, and the fungi Candida albicans and Candida glabrata. Due to difficulties in determining MIC values based on spectrophotometric methods, the antimicrobial activities of the ZnO, ZnO_c, ZnO_b and ZnO_i nanoparticles were assessed by determining the minimal bactericidal or fungicidal concentrations [62,63]. For this purpose, masses of nanoparticles ranging from 1.506 to 2.250 mg were weighted and poured on the wells of 6-well polystyrene plates. Overnight liquid cultures of bacteria and fungi were carried out by loop inoculating 25 mL of Mueller-Hinton broth (MHB) and Yeast Potato Dextrose (YPD) followed by incubation at 37 °C with orbital agitation (250 rpm). Overnight grown cultures were diluted 1:100 in MHB (bacteria) or Roswell Park Memorial Institute (RPMI) (fungi) and grown for additional 3 h at 37 °C with orbital agitation (250 rpm). Cultures were then adequately diluted in fresh media to obtain 5 × 10⁵ colony forming units (CFU) per mL. Adequate volumes of freshly inoculated MHB or RPMI broth, were added to the polystyrene plate wells to obtain nanoparticles final concentrations of 1000, 1250 or 1500 µg/ml. Plates were covered with aluminum foil to prevent light exposure and incubated for 24 h at 37 °C with agitation. After incubation, aliquots of 100 µl were spread on the surface of Luria Broth (LB, bacteria) solid media or Potato Dextrose Agar (PDA, fungi) and microbial growth was inspected after 24 h of plate incubation at 37 °C. The lowest nanoparticle concentration for which no colony forming units were detected was considered the minimal bactericidal or fungicidal concentration. Negative controls prepared with sterile culture media, and positive controls without nanoparticles were also carried out. Experiments were performed twice using triplicates per experiment.

4. Conclusions

In this work, a new class of halogenosilane-functionalized ZnO nanoparticles was successfully developed through a controlled chemical precipitation approach, demonstrating that surface modification represents an efficient strategy to tailor both physicochemical and biological properties of ZnO-based nanomaterials. Structural and morphological analyses confirmed that the incorporation of halogenosilanes leads to significant particle size reduction and improved dispersion, with bromopropylsilane-modified ZnO (ZnO_b) exhibiting the smallest particle size and most uniform morphology among the investigated nanomaterials.

Biological evaluation revealed a clear enhancement of anticancer activity for all halogenosilane-modified ZnO nanoparticles compared to unmodified ZnO, which can be attributed to improved cellular uptake and increased intracellular ROS generation. Interestingly, the relationship between cytotoxicity, apoptosis and ROS production suggests that multiple and potentially competing cell death pathways are involved, depending on nanoparticle surface characteristics. In particular, while unmodified ZnO appears to favour apoptosis-related mechanisms, quero systems, especially ZnO_b, may induce cell death through more complex ROS-driven processes.

Additional morphological evidence supporting these findings by SEM analysis of treated PC3 cells revealed distinct alterations in cell structure consistent with apoptosis and impaired cellular functionality. These observations reinforce the importance of nanoparticle–cell interactions in determining biological outcomes and further highlight the role of surface functionalization in modulating such interactions.

Overall, the present study demonstrates that halogenosilane functionalization enables modulation of the biological profile of ZnO nanoparticles, leading to enhanced anticancer activity together with distinct microbiocidal responses depending on the microbial system. These findings highlight the important role of surface chemistry in directing nanoparticle–cell interactions and support the possibility of tuning biological selectivity through rational surface modification.