All data are expressed as the mean of triplicates ± standard error (SE). The normality of the data distribution was confirmed using the Shapiro-Wilk test. Statistical significance was determined using one-way analysis of variance (ANOVA) followed by Tukey's post-hoc test for multiple comparisons. 95% confidence intervals (CI) are reported for key parameters. All analyses were performed using GraphPad Prism software version 5.0 (GraphPad Software, Inc., La Jolla, CA, USA).

The successful green synthesis of ZnO-NPs was observed as a visible change in the reaction mixture following the addition of L. sativum seed extract to the zinc nitrate solution (Fig. 2). The synthesis process involved the ionization of the precursor, Zn(NO₃)₂·6 H₂O, in an aqueous solution to produce Zn²⁺ ions. These ions were subsequently reduced by primary phytochemicals present in the L. sativum seed extract. The plant phytochemicals play a substantial role in nanoparticle biosynthesis. In this study, they facilitated the reduction of zinc ions (Zn²⁺) by acting as electron donors. Additionally, they functioned as capping agents, which was crucial for stabilizing the resulting ZnO-NPs. The L. sativum extract contains significant amounts of active compounds such as methylxanthines, flavonoids, phenolic acids, and saponins. These compounds are typically classified as antioxidants due to their ability to neutralize reactive oxygen species (ROS) and free radicals, as well as their capacity to chelate metals.

Therefore, these findings indicate that the antioxidants in the plant extract were responsible for the green synthesis of the nanoparticles. This is because such antioxidants can reduce metal ions and subsequently stabilize the newly formed nanoparticles.

The extract from L. sativum seeds and the biogenic zinc oxide nanoparticles (ZnO-NPs) were examined using UV-Vis spectroscopy to identify the surface plasmon resonance (SPR) of the nanoparticles (Fig. 3). The UV-Vis spectrum of the L. sativum extract exhibited two distinct peaks at 247 nm and 473 nm. In contrast, the spectrum of the biosynthesized ZnO-NPs displayed a broad absorption band ranging from 400 to 800 nm, with a prominent peak at 530 nm. This broad peak is characteristic of the SPR for metallic nanoparticles and is likely due to the aggregation of the biosynthesized ZnO-NPs. Additionally, the band observed at 247 nm in the ZnO-NPs spectrum may be attributed to the phytochemicals from the extract that functioned as reducing and capping agents.

The band gap energy of the ZnO-NPs was evaluated using the Tauc plot method and was found to be 3.08 eV (Fig. 4). This observed band gap energy aligns with a previous study that reported a similar band gap of 3.08 eV for ZnO-NPs synthesized using a green method with Musa acuminata peel extract. Similar findings for the band gap energy have also been reported in other studies.

TEM analysis revealed that the biosynthesized ZnO-NPs possessed a hexagonal structure (Fig. 5). The particle size distribution histogram indicated an average particle size of 32 nm (Fig. 6), a result which is consistent with those of prior reports. However, a previous study on the green synthesis of ZnO-NPs using Ocimum basilicum L. leaf extract reported a larger average particle size of 50 nm. Another report described sphere-like ZnO-NPs with an average size of 50 nm synthesized using Scutellaria baicalensis root extract. The smaller particle size (32 nm) achieved in this study, compared to other plant-mediated syntheses, can be attributed to the highly efficient capping and reducing action of the phytochemicals present in L. sativum extract. These compounds effectively control nucleation and growth during synthesis, resulting in smaller nanoparticles with a higher surface area-to-volume ratio, which enhances their biological activity.

Energy-dispersive X-ray (EDX) analysis was used to examine the biosynthesized ZnO-NPs and determine their elemental composition. The EDX spectrum revealed prominent peaks for zinc at energy levels of 1.1, 8.6, and 9.5 keV, which were identified as Zn Lα, Zn Kα, and Zn Kβ, respectively (Fig. 7). Additionally, a strong peak for oxygen was detected at an energy level of 0.5 keV (Table 1). Collectively, the elemental analysis confirmed the successful formation of ZnO nanoparticles. The mass percentages of elemental zinc and oxygen in the sample were determined to be 67.23% and 32.77%, respectively.

The crystalline nature of the biosynthesized ZnO-NPs was investigated using X-ray diffraction (XRD) analysis. The diffractogram demonstrated the presence of eleven distinct diffraction peaks at 2θ values of 31.82°, 34.65°, 36.36°, 47.82°, 56.73°, 62.98°, 66.49°, 68.00°, 69.23°, 72.64°, and 76.81° (Fig. 8). These peaks were indexed to the lattice planes (100), (002), (101), (102), (110), (103), (200), (112), (201), (004), and (202), respectively. This diffraction pattern is consistent with the standard pattern of JCPDS card no. 36-1451 and aligns with previous research, confirming the development of a hexagonal wurtzite crystal structure.

The average crystalline size of the ZnO-NPs was calculated using the Debye-Scherrer equation: D = Kλ/(β cosθ), where D is the average crystalline size, K is the Scherrer constant (0.94), λ is the X-ray wavelength (1.54178 Å), β is the full width at half maximum (FWHM) in radians, and θ is the Bragg diffraction angle. Based on the FWHM value of 0.2934 for the most intense peak (101) at 2θ = 36.36°, the average crystalline size was determined to be 29.78 nm.

The functional groups present in the L. sativum extract and on the surface of the biosynthesized ZnO-NPs were identified using FTIR analysis. The FTIR spectrum of the L. sativum extract revealed ten distinct absorption peaks at 3431.75, 2377.55, 2090.57, 1632.98, 1398.78, 1097.99, 1037.25, 899.40, 704.74, and 552.15 cm⁻¹. The spectrum of the biosynthesized ZnO-NPs showed vibrational frequencies at 3431.75, 3389.12, 2085.17, 1636.68, 1389.02, 1107.37, and 665.81 cm⁻¹ (Fig. 9). The broad band observed at 3431.75 cm⁻¹ in the extract spectrum was attributed to O-H stretching vibrations, Characteristic of phenolic compounds. The persistence of this band at 3431.75 cm⁻¹ and the appearance of a new band at 3389.12 cm⁻¹ in the ZnO-NP spectrum suggest that these phenolic compounds from the extract were involved in capping the nanoparticle surface. A weak band at 2377.55 cm⁻¹ in the extract, assigned to C ≡ C stretching of alkynes, was absent in the ZnO-NP spectrum. The band at 2090.57 cm⁻¹ in the extract spectrum shifted to a lower wavenumber (2085.17 cm⁻¹) in the ZnO-NP spectrum, which may indicate C-H stretching of aldehydes associated with the nanoparticles. Conversely, the band at 1632.98 cm⁻¹ (C = O stretching of amides) shifted to a higher wavenumber (1636.68 cm⁻¹). The band at 1398.78 cm⁻¹ (C-H stretching of alkanes) shifted to a lower wavenumber (1389.02 cm⁻¹). The peak at 1097.99 cm⁻¹ (C-O stretching of alcohols) shifted to a higher wavenumber (1107.37 cm⁻¹), affirming the interaction of these functional groups with the ZnO-NPs. Peaks in the extract at 1037.25, 899.40, and 704.74 cm⁻¹ were assigned to C-N stretching of amines, C = C bending of alkenes, and C-H bending of aromatic compounds, respectively; these were not present in the ZnO-NP spectrum, which suggests that the corresponding functional groups may have been involved in the reduction and stabilization processes. Finally, the peak observed at 552.15 cm⁻¹ in the extract shifted to 665.81 cm⁻¹ in the nanoparticle spectrum, a signature absorption band attributed to the Zn-O stretching vibration, confirming nanoparticle formation (Table 2).

The observed shifts in wavenumber and changes in band intensity between the L. sativum extract and the biosynthesized ZnO-NPs provide clear evidence of the functional groups involved in the reduction, capping, and stabilization processes. Specifically, the broadening and shift of the O-H stretching vibration suggest the involvement of phenolic compounds in reducing Zn²⁺ ions and subsequently capping the nanoparticle surface. Furthermore, shifts in the carbonyl (C = O) and amine (C-N) regions indicate that proteins and other organic molecules from the extract are also bound to the ZnO-NP surface, enhancing their stability and preventing aggregation. This correlation between the FTIR spectral data and the known phytochemical profile of L. sativum rich in polyphenols, flavonoids, and proteins strongly supports the role of these biomolecules in the green synthesis mechanism.

The surface charge and hydrodynamic diameter of the ZnO-NPs were assessed using a Zetasizer instrument based on photon correlation spectroscopy. The zeta potential measurements revealed a negative surface charge of approximately - 19.8 mV (Fig. 10). This negative potential is attributed to the capping by phytopolyphenols adsorbed onto the ZnO-NPs following the reduction process. Plant polyphenols are well-known to carry a negative charge, which they impart to the nanoparticle surface upon adsorption. These findings confirm the presence of a phytopolyphenol capping layer on the synthesized nanoparticles. Additionally, the biogenic ZnO-NPs exhibited a mean hydrodynamic diameter of 245.6 nm (Fig. 11).

Different concentrations of the biogenic ZnO-NPs showed antifungal activity against C. albicans and C. tropicalis strains (Fig. 12). The anticandidal activity of the ZnO-NPs was found to be higher against the C. tropicalis strain than against the C. albicans strain. The inhibition zone diameters for ZnO-NP concentrations of 0.125, 0.250, 0.500, and 1.0 mg/mL were 17.28 ± 0.51, 22.37 ± 0.48, 30.19 ± 0.32, and 38.63 ± 0.29 mm, respectively, against the C. tropicalis strain. However, the inhibition zone diameters for ZnO-NP concentrations of 0.125, 0.250, 0.500, and 1.0 mg/mL were 10.98 ± 0.56, 11.76 ± 0.24, 13.58 ± 0.43, and 18.69 ± 0.34 mm against the C. albicans strain, respectively (Table 3). Previous investigation demonstrated that the biosynthesized ZnO nanoparticles demonstrated potent antimicrobial efficacy against key pathogens including E. coli, E. faecalis, S. aureus, and S. mutans, with zones of inhibition up to 20.0 mm, MIC values of 15 to 25 µg/mL, and complete bactericidal activity within 8 to 12 h, while also exhibiting high biocompatibility in zebrafish at doses up to 1 mg/mL . The MICs of the biosynthesized ZnO-NPs were found to be 62.5 and 125 µg/mL against C. tropicalis and C. albicans strains, respectively. The synergistic activity of the MIC concentrations of the biosynthesized ZnO-NPs with common antifungal agents was evaluated using the standard disk diffusion method. The highest synergistic activity of the ZnO-NPs was detected with nystatin against C. albicans and C. tropicalis strains with IFA values of 0.85 and 0.99, respectively (Table 4). The combined effect of ZnO-NPs with nystatin was significantly higher than that of nystatin alone, with p-values of 0.0017 and 0.0096, respectively. Moreover, significant additive effects were observed for ZnO-NPs with both fluconazole (IFA = 0.62, p = 0.0426) and terbinafine (IFA = 0.77, p = 0.0031) against the C. albicans strain. The combination with ketoconazole against C. albicans showed an IFA of 0.47 but was not statistically significant (p > 0.05). Against C. tropicalis strain, significant additive effects were found for ZnO-NPs with ketoconazole (IFA = 0.78, p = 0.0316) and fluconazole (IFA = 0.79, p = 0.0011). The combination with amphotericin B (IFA = 0.58) was not significant. In contrast, the combined effect of ZnO-NPs with itraconazole, terbinafine, and amphotericin B against C. tropicalis strain was not significantly different compared to the antifungals alone (p > 0.05). Furthermore, no significant difference was found for ZnO-NPs combined with ketoconazole, itraconazole, and amphotericin B against C. albicans strain (p > 0.05).

The FICI results quantitatively confirmed the interactions that were detected by the disk diffusion assay. The most potent synergistic effects were observed for the combination of ZnONPs with nystatin against both C. albicans (FICI = 0.38) and C. tropicalis (FICI = 0.25). Additive interactions were confirmed for ZnONPs with fluconazole and terbinafine against C. albicans (FICI = 0.75 and 0.63), and with ketoconazole and fluconazole against C. tropicalis (FICI = 0.75 and 0.63). For other combinations, such as those with itraconazole and amphotericin B against both species, the indifferent FICI scores aligned with their low, non-significant IFA values. Crucially, the FICI score formally identified an antagonistic interaction for the combination of terbinafine and ZnONPs against C. tropicalis (FICI = 4.00).

The antimicrobial effects of ZnO-NPs are attributed to multiple mechanisms, including the production of reactive oxygen species (ROS)compromising the integrity of cell membranesreleasing toxic zinc ions (Zn²⁺), interacting with microbial DNA and proteinsand triggering oxidative stress. Zinc oxide nanoparticles can produce ROS like hydroxyl radicals (*OH), hydrogen peroxide (H₂O₂), and superoxide anions (O₂*-) when exposed to light or specific conditions. These ROS are extremely reactive and can cause damage to microbial cell membranes, lipids, proteins, and DNA, ultimately resulting in cell death. Moreover, ZnO-NPs can directly interact with microbial cell membranes, leading to membrane damage. This disturbance can enhance membrane permeability, causing the leakage of cellular materials and, ultimately, cell death and lysis. The biosynthesized ZnO-NPs can break down and release zinc ions (Zn²⁺) that bind to microbial proteins and enzymes, disrupting their normal functions and hindering cell growth. Additionally, Zn²⁺ can interfere with crucial metal ion transport systems, adding further stress to the candidal cells. Due to their small size, ZnO-NPs can infiltrate microbial cells and interact with internal components such as DNA and proteins. This interaction may result in DNA damage, proteins losing their structure, and enzymes becoming inactive, ultimately disrupting essential cellular functions.

The notable difference in synergy between nystatin and amphotericin B (AmB), both polyene antifungals, can be attributed to their distinct physicochemical properties and mechanisms. While both target ergosterol, nystatin molecules are smaller and form smaller, more transient pores in the fungal membrane compared to the larger, more stable channels formed by AmB aggregates. We hypothesized that the smaller nystatin-induced pores are sufficient to cause membrane leakage but are also more permissive for the subsequent influx of ZnO-NPs or Zn²⁺ ions, leading to amplified intracellular oxidative damage.

The profound synergy observed between ZnO-NPs and nystatin, particularly against C. tropicalis (IFA: 0.99), is of significant clinical relevance. By combining ZnO-NPs with a conventional antifungal, the effective dose of the drug required to achieve fungicidal activity can be substantially lowered. This approach has a high potential to reduce the drug concentration below its toxic threshold, thereby mitigating dose-limiting side effects like the nephrotoxicity associated with high doses of polyenes. This is a crucial strategy for managing chronic or recurrent infections, especially in debilitated patients.

This synergy is likely multimodal: nystatin binds to ergosterol, creating pores in the fungal membranewhich potentially facilitates the increased uptake of ZnO-NPs (Fig. 13). Subsequently, the internalized NPs can induce massive ROS generation and release Zn²⁺ ionsleading to catastrophic oxidative damage and disruption of metabolic enzymes. The weaker synergy with amphotericin B, another polyene, might be attributed to differences in their binding affinity to ergosterol or their aggregation state in solution. The synergy with azoles like fluconazole may stem from the combined stress of ergosterol biosynthesis inhibition (azole) and direct membrane/oxidative damage (ZnO-NPs).

The antioxidant activity of the biogenic ZnO NPs synthesized using L. sativum seeds extract was assessed by DPPH scavenging assay. The DPPH inhibition percentages were found to be 28.14 ± 0.87, 36.48 ± 1.14, 48.32 ± 1.35, 61.87 ± 1.56, 78.95 ± 1.64 and 86.78 ± 1.83 for the different ZnONPs concentrations of 50, 100, 200, 400, 800 and 1600 µg/mL, respectively (Fig. 14). Accordingly, the antioxidant activity of ZnO-NPs was concentration dependent. DPPH is a well-known and stable synthetic solid radical commonly used to assess the antioxidant potential of various compounds. The spectrophotometer was used to quantify the reduction of DPPH by receiving hydrogen or electrons from ZnO nanoparticles, which caused the colour to change from purple to yellow. Linear regression analysis revealed that the IC of ZnO NPs was 335.48 µg/mL whereas the IC of ascorbic acid was 131.03 µg/mL. Collectively, the biosynthesized ZnO-NPs have potential for creating potent antioxidants that could be utilized in treating numerous diseases linked to oxidative stress.

The antiproliferative activity of the biosynthesized ZnONPs was evaluated using MTT assay against HUH7 and WI38 cell lines. The cytotoxic effect of ZnONPs was found to be concentration dependent against HUH7 cell line, demonstrating relative cell viability percentages of 78.58, 64.85, 58.73 and 37.85% at the concentrations of 25, 50, 100 and 200 µg/ml, respectively. On the other hand, the bioinspired ZnONPs revealed a lower cytotoxic effect against WI38 normal cell line demonstrating relative cell viability percentages of 62.96 and 45.92% at ZnONPs concentrations of 100 and 200 µg/ml, respectively. As shown in Fig. 15, the cytotoxic effect of ZnONPs was concentration-dependent in both cell lines. Post-hoc analysis confirmed that ZnONPs exhibited significantly greater cytotoxicity in HUH-7 cancer cells compared to WI-38 normal cells at concentrations of 100 µg/ml (p < 0.01) and 200 µg/ml (p < 0.001). Specifically, at 100 µg/ml, viability was reduced to 58.73 ± 6.62% in HUH-7 cells versus 62.96 ± 5.92% in WI-38 cells. At 200 µg/ml, viability was further reduced to 37.85 ± 5.41% in HUH-7 cells, which was significantly lower than the 45.92 ± 3.12% observed in WI-38 cells. The half-maximal inhibitory concentration (IC₅₀) was determined by fitting the cell viability data to a four-parameter logistic (4PL) non-linear regression model. The analysis revealed IC₅₀ values of 145.2 µg/mL and 237.6 µg/mL for the HUH7 and WI38 cell lines, respectively. Zinc oxide nanoparticles exert their selective anticancer effects primarily through the induction of reactive oxygen species (ROS)-mediated oxidative stress and apoptosis pathways. Upon cellular internalization, ZnONPs generate excessive ROS, including superoxide anions and hydroxyl radicals, which overwhelm the antioxidant defense systems of cancer cells, leading to mitochondrial membrane potential disruption, cytochrome c release, and activation of caspase cascades (caspase-3 and caspase-9). This oxidative stress is further amplified by the release of Zn²⁺ ions due to the acidic tumor microenvironment, which promotes ZnO NP dissolution and enhances intracellular zinc concentrations, causing additional DNA damage and cell cycle arrest. The selective toxicity toward cancer cells is attributed to their higher metabolic activity and elevated basal ROS levels, making them more vulnerable to oxidative damage compared to normal cells. Recent studies also highlight the role of ZnONPs in inhibiting angiogenesis and epithelial-mesenchymal transition (EMT), thereby reducing metastasis and chemotherapy resistance in aggressive cancers.

The toxicity and efficacy of ZnONPs are profoundly influenced by their physicochemical properties, including size, shape, surface charge, and functionalization. The superior cytotoxic effect of the smaller nanoparticles (≤ 50 nm), such as the 32 nm particles in our study, stems from their higher surface area-to-volume ratio, which promotes increased cellular uptake, elevated ROS generation, and accelerated ion release rates. Moreover, surface functionalization with bioactive compounds (e.g., phenolic compounds from plant extracts) further modulates stability, dispersibility, and cellular interactions. Moreover, hexagonal and spherical morphologies promote membrane disruption and internalization, while anisotropic structures like nanorods induce mechanical damage to cellular structures. The green synthesis approach of ZnONPs using plant extracts as L. sativum extract not only reduces ecological toxicity but also enhances anticancer efficacy by capping phytochemicals that synergize with ZnONPs to target cancer-specific pathways.

The selectivity index (SI), a crucial parameter for evaluating therapeutic safety, is defined as the ratio of IC₅₀ (normal cells) to IC₅₀ (cancer cells). Here, the SI for HUH7 cells versus WI38 cells is ~ 1.64 (237.6/145.2), indicating a clear and favorable selectivity for cancer cells. More importantly, the MIC values for Candida spp. (62.5-125 µg/mL) are significantly lower than the IC₅₀ for normal WI38 cells (237.6 µg/mL), indicating a strong safety margin for antimicrobial applications. This compelling differential toxicity suggests the high potential biosafety and therapeutic utility of the biosynthesized ZnONPs.

The implications of this study are particularly significant for disability patients. The demonstrated synergy between biogenic ZnO NPs and conventional antifungals, especially nystatin, offers a promising avenue to combat drug-resistant candidiasis, a common and serious threat in this population. By significantly enhancing the efficacy of existing drugs, this approach allows for the use of lower doses of antifungals to achieve a therapeutic effect. This dose-reduction strategy has a direct clinical benefit for disability patients as it can potentially mitigate the severe dose-limiting side effects, such as nephrotoxicity associated with amphotericin B or hepatotoxicity with azoles, which are especially dangerous for individuals with pre-existing organ dysfunction or polypharmacy. Consequently, this nano-adjuvant therapy could lead to more successful treatment outcomes, reduce the frequency of recurrent infections, decrease hospitalization periods, and ultimately improve the quality of life and long-term health prospects for this vulnerable patient group. However, it is crucial to address the limitations of this work, which include the necessity to perform in vivo studies to validate these findings and the absence of biofilm disruption assays, a key virulence factor in persistent infections.