Introduction

Nanotechnology is an emerging scientific and technological topic that is gaining attention from several sectors. Metallic nanoparticles (NPs) are often used in the biomedical sector1. The reactivity of NPs is much greater than that of bulk materials due to their extensive surface area2. Biologists are becoming increasingly interested in the utilization of NPs for various applications, including drug delivery, gene delivery, and material detection in biological samples3. Moreover, NPs also show potential in combating pathogens and cancers4. A previous investigation demonstrated the potential biological activities of biologically synthesized NPs, such as AuNPs synthesized using fresh leaves of Polygala elongata, which demonstrated potential anticancer activity against the lung cancer cell line (A549)5. Moreover, fluorinated graphene oxide nanosheets derived from Lissachatina fulica snail mucus demonstrated antibacterial efficiency against bacterial pathogens and anticancer activity against the pancreatic cancer cell line (PANC1) at low concentrations6. Another study reported that an Acorus calamus-mediated CuFe2O4/reduced graphene oxide nanocomposite showed antibacterial activity, demonstrating relative inhibition zone diameters ranging from 10 to 13 mm against the tested strains, and also low cytotoxic activity against the MCF-7 cell line with an IC50 of 360 µg/mL7. Similarly, a ZnO/g-C3N4/V2O5 heterogeneous nanocomposite revealed potential antibacterial and photocatalytic activities8. Zinc oxide nanoparticles (ZnO-NPs) offer a wide range of capabilities that make them useful in several disciplines, including chemical, physical, and biological applications9. The semiconducting and oxidizing properties of ZnO nanoparticles play an important role in their biological efficiency. The size and crystalline structure of ZnO nanoparticles determine their reactivity and chemical properties10. Furthermore, ZnO-NPs have substantial photocatalytic activity on several substances, including dyes, medicines, and toxins11. Studies have shown that ZnO-NPs are more compatible with the physiological environments of the body compared to their bulk form. This suggests that their medicinal properties may provide distinct advantages12. Green synthesis is a commonly used technique utilized by biological systems, including microorganisms, plant extracts, or their constituents, for the formation of nanoparticles13. Generally, biological-based synthesis is more advantageous than alternative chemical and physical approaches in terms of cost-effectiveness, single-step processing, and absence of chemical waste throughout the process14. Therefore, the synthesis of nanoparticles using biological means is a secure and non-toxic procedure suitable for pharmaceutical and medical applications15. The production of bio-based nanoparticles for use in the pharmaceutical, cosmetic, culinary, and environmental industries has seen significant growth in recent decades16. A previous investigation demonstrated that biogenic ZnONPs synthesized using Pleurotus sajor-caju extract demonstrated antibacterial activity against Staphylococcus aureus (6.2 ± 0.1 mm), Streptococcus mutans (5.4 ± 0.4 mm), and Bacillus subtilis (5.2 ± 0.1 mm)17. Moreover, Hypsizygus ulmarius-derived ZnO nanoparticles revealed potential antibacterial activity with relative inhibition zone diameters of 9.0 ± 0.1 mm and 14.2 ± 0.2 mm against K. pneumoniae and S. aureus, respectively, and also potent anticancer activity against MCF-7 breast cancer cells with relative LC50 values of 29.07 µg/mL18. Zinc oxide nanoparticles are very promising nanomaterials that have been shown to possess excellent physiochemical stability and unique surface attraction capabilities. As a result, they may be effectively used for many biomedical applications, including antifungal and antibacterial treatments19. Moreover, ZnO NPs have been acknowledged by the FDA-USA as the safest kind of nanoparticles20. Multiple investigations have been conducted to assess the fungicidal effects of ZnO NPs on various fungal pathogens21,22. These studies have shown that ZnO NPs exhibit little toxicity towards human cells23. The toxicity mechanisms of ZnO-NPs towards bacteria or fungi mostly rely on the size, shape, and concentration of ZnO-NPs, as well as the specific kind of medium used. In general, as the size of a particle is smaller, its surface area to volume ratio increases, resulting in stronger antimicrobial actions24. The physical interaction between ZnO-NPs and the fungal cell wall results in the disruption of the cell wall’s structure25. This interaction also triggers the excessive creation of reactive oxygen species (ROS), including hydroxyl groups, superoxide anion radicals, and hydrogen peroxide, inside the cells. Consequently, these ROS may cause the demise of the cells. The amount of ROS produced is directly correlated with the extent of the organism’s surface area that is in contact with the ZnO NPs. In addition, several studies have shown that ZnO-NPs have dose-dependent effects, wherein larger concentrations of ZnO-NPs result in increased antifungal activity26. Candida spp. are the predominant fungal pathogens that opportunistically infect people. They are often acquired in hospitals and rank as the fourth most prevalent cause of nosocomial infections27. C. albicans is the primary factor responsible for almost all forms of Candidiasis28. Conventional antifungal medications work by either suppressing the development of fungal cells or causing their death. Both scenarios involve medications that impose significant selection pressure and may readily result in the development of drug resistance29. Moreover, the challenge of managing candidiasis is profoundly magnified in individuals with disabilities. This population often experiences compromised immune function, prolonged hospitalization, frequent use of broad-spectrum antibiotics, and the presence of medical devices like catheters and ventilators, all of which are significant risk factors for fungal colonization and subsequent infection. Furthermore, the high prevalence of drug-resistant Candida strains, such as fluconazole-resistant C. albicans creates a critical therapeutic impasse. Recurrent and persistent infections in this vulnerable demographic lead to increased morbidity, extended rehabilitation times, and a substantial additional burden on both patients and healthcare systems, underscoring an urgent need for novel, effective, and safer antifungal strategies30. Recently, researchers have focused on targeting fungal virulence as a potential therapeutic target31. Suppressing the conversion of C. albicans into its disease-causing state by reducing the virulence of the fungus has emerged as a successful approach for combating fungal infections32. A previous investigation demonstrated that biogenic ZnO NPs from L. sativum seeds extract exhibited strong antimicrobial activity, with higher efficacy against S. aureus (23 ± 1.25 mm inhibition at 120 mg/mL) than E. coli (16 ± 1.00 mm), while remaining hemocompatible at low doses33. Another previous investigation demonstrated that biogenic ZnONPs formulated using L. sativum seed extract exhibited broad-spectrum antibacterial activity against both Gram-positive (S. aureus: 18 ± 1.1 mm, Micrococcus luteus: 15 ± 1.2 mm) and Gram-negative pathogens (Salmonella enterica serovar Typhi: 15 ± 1.2 mm, Salmonella Setubal: 18 ± 1.4 mm, Enterobacter aerogenes: 14 ± 1 mm) at 4 mg/mL. The NPs showed comparable efficacy to the antibiotic cefexime (e.g., S. aureus: NPs 18 mm vs. cefexime 19 mm), while the crude plant extract alone displayed no significant inhibition (< 12 mm). These findings highlight the potential of L. sativum-derived ZnO NPs as a promising antimicrobial agent34. The antimicrobial activity of biogenic ZnONPs was assessed via the Kirby-Bauer disk diffusion method, revealing significant antibacterial effects (28.74 ± 1.74 mm for S. aureus, 21.04 ± 0.34 mm for E. coli) and moderate antifungal activity (26.11 ± 0.86 mm for C. tropicalis, 18.76 ± 1.00 mm for C. albicans)35. A previous work presented an inexpensive and eco-friendly synthesis of spherical, 5–35.5 nm ZnO NPs using Ocimum tenuiflorum, which demonstrated potent antibacterial and antibiofilm activity against key gram-positive and gram-negative pathogens while exhibiting low toxicity in a zebrafish model36.

While the standalone antifungal activity of ZnO-NPs is established, their potential to synergize with conventional drugs to overcome resistance is a burgeoning area of research. Recent studies have shown promising synergistic effects between biogenic silver nanoparticles and azole antifungal agents such as fluconazole against resistant clinical isolates of six Candida species37.

This combination therapy approach can resensitize resistant strains and reduce the required dosage of toxic antifungals, mitigating side effects and delaying resistance development. Collectively, the previous investigations on L. sativum-derived ZnO NPs focused on antibacterial activity, while the current study investigated the antifungal activity of the biogenic ZnONPs from L. sativum seeds against candidal pathogens for the first time. Moreover, the synergistic activity of ZnONPs with conventional antifungal agents such as Ketoconazole, Fluconazole, Itraconazole, Nystatin, Terbinafine, and Amphotericin B was evaluated against candidal pathogens. Furthermore, the study investigated the antioxidant and selective cytotoxic properties, thereby presenting a multifaceted biomedical application.

Materials and methods

Chemicals and reagents

All chemicals and solvents utilized in this study were of analytical grade and were used without further purification. Zinc nitrate hexahydrate (Zn(NO₃)₂·6 H₂O, ≥ 99%) and sodium hydroxide (NaOH, pellets, ≥ 97%) were procured from Sigma-Aldrich (USA) for the synthesis of zinc oxide nanoparticles. For the antifungal assays, standard antifungal drugs including ketoconazole, fluconazole, itraconazole, nystatin, terbinafine, and amphotericin B were obtained from Himedia (India). The antioxidant activity was assessed using 1,1-diphenyl-2-picrylhydrazyl (DPPH, ≥ 95%, Sigma-Aldrich, USA). For the cytotoxicity assay, methylthiazolyl diphenyl-tetrazolium bromide (MTT, ≥ 98%) and dimethyl sulfoxide (DMSO, ≥ 99.9%) were purchased from Sigma-Aldrich (USA). High-purity methanol (CH₃OH, 99.8%) and absolute ethanol (C₂H₅OH, 99.9%) were used as solvents for sample preparation and washing procedures. All aqueous solutions were prepared using distilled water. Sabouraud Dextrose Agar (SDA) and Roswell Park Memorial Institute (RPMI-1640) cell culture medium were acquired from Himedia (India) and Millipore Sigma (USA), respectively. Fetal bovine serum (FBS) and phosphate-buffered saline (PBS) tablets were sourced from Thermo Fisher Scientific (USA).

Preparation of L. sativum seed extract

The selection of L. sativum seed extract for the green synthesis of ZnO-NPs was predicated on its established high concentration of bioactive reducing and stabilizing agents, particularly polyphenolic compounds such as rosmarinic, caffeic, cinnamic, p-coumaric, carnosic, and ferulic acid38. These bioactive compounds are highly efficacious reducing agents; their phenolic hydroxyl and other functional groups donate electrons to reduce metal salt precursors (Zn²⁺) to zerovalent metal atoms, which subsequently oxidize to form metal oxide nanoparticles39. Furthermore, these natural compounds act as effective capping and stabilizing agents, preventing nanoparticle aggregation and ensuring colloidal stability. Figure 1 shows the diagrammatic illustration of the synthesis of ZnO nanoparticles using phytoconstituents from L. sativum seeds extract.

The L. sativum seeds were purchased from a local market in Riyadh, Saudi Arabia. The collected seeds were taxonomically identified at the Herbarium of the Department of Botany and Microbiology, College of Science at King Saud University. The plant material was deposited in the Herbarium with voucher number of KSU-202,506. No specific permissions were required for the acquisition of these commercially available seeds. The aqueous extract was prepared by adding 50 g of powdered plant seeds to a 500 mL flask containing 200 mL of distilled water. The flask was placed on a magnetic stirrer hot plate and stirred for a duration of 2 h at a temperature of 60 °C. The extract was filtered twice using a nylon cloth to eliminate solid plant residues. This was followed by further filtration using Whatman filter paper for a total of three filtrations to ensure the removal of any remaining residues. Subsequently, the extract was sterilized by filtration through a 0.45 μm Millipore membrane filter. Finally, the resulting extract was stored in a refrigerator at a temperature of 4 °C for subsequent use40. All methods were performed in accordance with the relevant guidelines and regulations.

Fig. 1
figure 1

Schematic diagram of the green synthesis of ZnO NPs using L. sativum seed extract, where phytochemicals act as both reducing and capping agents.

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Green synthesis of ZnO-NPs

A 95 mL volume of a 0.01 M aqueous solution of zinc nitrate hexahydrate was prepared. To this solution, 5 mL of L. sativum seed extract was added dropwise under constant stirring. The pH of the reaction mixture was monitored using a calibrated digital pH meter and adjusted to 12.0 by the slow addition of a 0.02 M NaOH solution. The mixture was then continuously agitated using a magnetic stirrer at 500 rpm for 2 h at 60 °C under ambient light conditions. The formation of a white precipitate indicated the synthesis of ZnO-NPs. The precipitate was collected by centrifugation at 10,000 rpm for 20 min. The resulting pellet was washed twice with distilled H₂O and three times with absolute ethanol to remove impurities. Finally, the sample was dried in an oven at 60 °C for 24 h. Subsequently, the dehydrated ZnO-NPs were calcined in a muffle furnace at 400 °C for 2 h41.

Characterization (UV-Vis, TEM, EDX, XRD, FTIR, DLS/Zeta)

Various Characterization techniques were employed to investigate the Chemical and physical properties of the synthesized ZnO-NPs. The biosynthesis was initially verified using a UV-Vis spectrophotometer by measuring absorbance across wavelengths from 200 to 800 nm. Transmission electron microscopy (TEM) was performed to analyze the particle morphology and size. The elemental composition (mass percentages of zinc and oxygen) was determined using a scanning electron microscope (SEM) equipped with an energy dispersive X-ray (EDX) analyzer (JEOL, JSM-6380 LA, Tokyo, Japan). The crystalline structure of the thermally annealed sample was analyzed using an X-ray diffraction (XRD) instrument (Shimadzu XRD-6000 diffractometer, Shimadzu, Columbia, USA) equipped with a Cu radiation source operating at 45 kV and 40 mA. The average crystalline size was calculated using the Scherrer equation. Functional groups present on the nanoparticle surface were identified by Fourier-transform infrared (FTIR) spectroscopy, with scans performed in the range of 400–4000 cm⁻¹. Finally, the zeta potential, hydrodynamic size distribution, and polydispersity index (PDI) were assessed using a dynamic light scattering (DLS) system (Malvern Zetasizer Nano).

Antifungal activity of different concentrations of the biosynthesized ZnO-NPs

The antifungal activity of various concentrations (0.125, 0.250, 0.500, and 1.000 mg/disk) of ZnO-NPs was evaluated against two candidal strains, C. albicans (ATCC 29213) and C. tropicalis (ATCC 33592), using the standard disk diffusion assay. Fungal suspensions were prepared in 0.9% sodium Chloride, and the turbidity was adjusted to a 0.5 McFarland standard. Fresh Sabouraud dextrose agar (SDA) plates were swabbed uniformly with 100 µL of each microbial suspension. The dried ZnO-NPs were dissolved in methanol to create a 10 mg/mL stock solution, which was Sonicated for 30 min to ensure uniform dispersion. Sterile filter paper disks (8 mm diameter) were impregnated with the stock solution to achieve the desired final concentrations. Disks impregnated with methanol alone served as a negative control, and disks containing 30 µg of terbinafine served as a positive control. All plates were incubated at 35 °C for 48 h, after which the diameters of the zones of inhibition were measured using a Vernier caliper42.

The minimum inhibitory concentrations (MICs) were determined using a broth microdilution assay in 96-well microtiter plates43. Subsequently, the MIC and the next two higher concentrations were streaked onto SDA plates to determine the minimum fungicidal concentrations (MFCs). The plates were incubated at 35 °C for 48 h, and the lowest concentration that showed no visible fungal growth was recorded as the MFC.

Synergistic antifungal effectiveness of ZnO-NPs with antifungal agents

The combined antifungal activity of ZnO-NPs with standard antifungal drugs, ketoconazole (30 µg), fluconazole (25 µg), itraconazole (10 µg), nystatin (100 µg), terbinafine (30 µg), and amphotericin B (100 µg), was evaluated using the disk diffusion assay. These antifungals, representing distinct classes (polyenes, azoles, and an allylamine) with differing mechanisms of action, were selected to assess broad-spectrum synergy. Filter paper disks were impregnated with the stated concentrations of the antifungal agents alone and in combination with the MIC value of the ZnO-NPs to evaluate the combined effect. Negative (methanol) and positive (antifungal alone) control disks were also included. The loaded disks were placed on seeded SDA plates, prepared as described in Sect. 2.5. After a 48-hour incubation period, the zones of suppression were measured. The increase in fold area (IFA) was then calculated using the following equation:

$$\:\mathbf{I}\mathbf{F}\mathbf{A}\:=\:(\mathbf{b}^2\:\ - \:\mathbf{a}^2)\:/\:\mathbf{a}^2$$

where a represents the zone of inhibition for the antifungal agent alone, and b represents the inhibition zone for the combination of the antifungal agent and ZnO-NPs44. Moreover, the fractional inhibitory concentration (FIC) index was estimated using the formula described in previous studies45,46.

\(\:\text{F}\text{I}\text{C}\:\text{i}\text{n}\text{d}\text{e}\text{x}\:\left(\text{F}\text{I}\text{C}\text{I}\right)\:=\frac{\text{M}\text{I}\text{C}\:\text{o}\text{f}\:\text{d}\text{r}\text{u}\text{g}\:\text{A}\:\text{i}\text{n}\:\text{t}\text{h}\text{e}\:\text{c}\text{o}\text{m}\text{b}\text{i}\text{n}\text{a}\text{t}\text{i}\text{o}\text{n}}{\text{M}\text{I}\text{C}\:\left(\text{A}\right)}+\frac{\text{M}\text{I}\text{C}\:\text{o}\text{f}\:\text{d}\text{r}\text{u}\text{g}\:\text{B}\:\text{i}\text{n}\:\text{t}\text{h}\text{e}\:\text{c}\text{o}\text{m}\text{b}\text{i}\text{n}\text{a}\text{t}\text{i}\text{o}\text{n}}{\text{M}\text{I}\text{C}\:\left(\text{B}\right)}\), whereas A referred to ZnO NPs and B referred to antifungal agents. The FICI detected as follows: FICI ≤ 0.5 synergistic effect, FICI = 0.5 to 1 additive effect, 2 ≤ FICI < 4 indifferent, and 4 < FICI antagonism.

Antioxidant activity

The antioxidant activity of various concentrations of ZnO-NPs (50, 100, 200, 400, 800, and 1600 µg/mL) was evaluated using the 1,1-diphenyl-2-picrylhydrazyl (DPPH) radical scavenging assay. Briefly, 2 mL of a 1 mM DPPH solution was mixed with each concentration of ZnO-NPs. The solutions were incubated in the dark for 30 min. The absorbance was then measured at 517 nm using a UV-Vis spectrophotometer. The percentage of DPPH scavenging activity was calculated using the following formula:

% DPPH scavenging = [(A − B)/A] × 100, where A is the absorbance of the standard and B is the absorbance of the sample47.

Cytotoxicity assay

The cytotoxicity of the biosynthesized ZnO-NPs was evaluated on human hepatoma (HUH7) and normal lung fibroblast (WI38, ATCC CCL-75) cell lines using the methylthiazolyl diphenyl-tetrazolium bromide (MTT) assay. The HUH7 cell line was purchased from Millipore Sigma (USA), and the WI38 cell line was kindly donated by the Department of Zoology, College of Science, King Saud University, Saudi Arabia. Cells used in experiments were between passages 15–25 and were routinely confirmed to be free of mycoplasma contamination.

Cells were seeded in a 96-well tissue culture plate at a density of 1 × 10⁵ cells/mL in 100 µL of growth medium per well and incubated at 37 °C for 24 h. After removing the growth medium, the cells were treated with 0.1 mL of serially diluted ZnO-NPs in RPMI media containing 2% serum. Control wells were treated with serum-containing media only. The plate was incubated at 37 °C for 48 h. Thereafter, 10 µL of a 5 mg/mL MTT solution in PBS was added to each well. The plate was incubated for 4 h at 37 °C with 5% CO₂ to allow formazan crystal formation. The formazan crystals were then dissolved by adding 200 µL of dimethyl sulfoxide (DMSO) to each well with gentle agitation for 5 min. The absorbance was measured at a wavelength of 560 nm. The half-maximal inhibitory concentration (IC₅₀) was calculated by fitting cell viability data to a sigmoidal dose-response model using non-linear regression48.

Statistical analysis

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).

Results and discussion

Green synthesis of ZnO-NPs

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 extract49. The plant phytochemicals play a substantial role in nanoparticle biosynthesis50. 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-NPs51. The L. sativum extract contains significant amounts of active compounds such as methylxanthines, flavonoids, phenolic acids, and saponins52. 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 metals53.

Therefore, these findings indicate that the antioxidants in the plant extract were responsible for the green synthesis of the nanoparticles54. This is because such antioxidants can reduce metal ions and subsequently stabilize the newly formed nanoparticles55.

Fig. 2
figure 2

Color change observed during the green synthesis of ZnO NPs. The transition upon addition of L. sativum seeds extract to the zinc precursor solution indicates nanoparticle formation.

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Spectral analysis of ZnO-NPs

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 nm56. This broad peak is characteristic of the SPR for metallic nanoparticles and is likely due to the aggregation of the biosynthesized ZnO-NPs57. 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 agents58.

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 extract59. Similar findings for the band gap energy have also been reported in other studies60,61.

Fig. 3
figure 3

UV-Vis spectra of L. sativum extract (black) and biosynthesized ZnO NPs (red). ZnO NP spectrum shows a broad plasmon resonance peak at 530 nm.

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Fig. 4
figure 4

Tauc plot for band gap determination of ZnO NPs. The calculated optical band gap is 3.08 eV.

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TEM analysis of ZnO-NPs

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 reports62,63,64. 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 nm65. Another report described sphere-like ZnO-NPs with an average size of 50 nm synthesized using Scutellaria baicalensis root extract66. 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.

Fig. 5
figure 5

TEM micrograph of the biosynthesized ZnO-NPs showing hexagonal morphology. Scale bar: [100] nm.

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Fig. 6
figure 6

Particle size distribution histogram from TEM analysis. Average particle size is 32 nm.

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EDX analysis of the biogenic ZnO-NPs

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.

Fig. 7
figure 7

EDX spectrum of ZnO NPs confirming composition, with characteristic Zn and O peaks.

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Table 1 Elemental map of the biogenic ZnONPs.
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XRD analysis of ZnO-NPs

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 structure67,68.

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.

Fig. 8
figure 8

XRD pattern of ZnO NPs. All diffraction peaks are indexed to the hexagonal wurtzite structure of ZnO (JCPDS No. 36-1451).

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FTIR analysis of ZnO-NPs

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 surface69. A weak band at 2377.55 cm⁻¹ in the extract, assigned to C ≡ C stretching of alkynes, was absent in the ZnO-NP spectrum70. 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)71. 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-NPs72. 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 processes73,74,75. 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)76.

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.

Fig. 9
figure 9

FTIR spectra of L. sativum extract (red) and ZnO NPs (black). Peak shifts indicate phytochemicals acting as reducing and capping agents.

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Table 2 Functional groups of L. sativum seeds extract and the biosynthesized ZnO-NPs.
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Zeta potential analysis of ZnONPs

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 nanoparticles77. Additionally, the biogenic ZnO-NPs exhibited a mean hydrodynamic diameter of 245.6 nm (Fig. 11).

Fig. 10
figure 10

Zeta potential distribution of ZnO NPs. The average value (−19.8 mV) indicates moderate colloidal stability.

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Fig. 11
figure 11

Hydrodynamic size distribution of ZnO NPs in suspension by DLS. The average diameter is 245.6 nm.

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Antifungal activity of different concentrations of the bioprepared ZnO-NPs

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 78. 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)78compromising the integrity of cell membranes79releasing toxic zinc ions (Zn²⁺)80, interacting with microbial DNA and proteins81and triggering oxidative stress82. Zinc oxide nanoparticles can produce ROS like hydroxyl radicals (•OH), hydrogen peroxide (H₂O₂), and superoxide anions (O₂•−) when exposed to light or specific conditions83. These ROS are extremely reactive and can cause damage to microbial cell membranes, lipids, proteins, and DNA, ultimately resulting in cell death84. 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 lysis85. 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 cells86. 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 functions87.

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 aggregates88. 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 polyenes89. 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 membrane90which potentially facilitates the increased uptake of ZnO-NPs (Fig. 13). Subsequently, the internalized NPs can induce massive ROS generation91 and release Zn²⁺ ions92leading to catastrophic oxidative damage and disruption of metabolic enzymes93. 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)94.

Fig. 12
figure 12

Antifungal activity of ZnO NPs against C. albicans and C. tropicalis. Zones of inhibition for various NP concentrations (0.125-1.000 mg/disk), with terbinafine (30 µg/disk) as positive control and methanol solvent as negative control disks.

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Table 3 Antifungal activity of the biosynthesized znonps. Against C. albicans and C. tropicalis strains.
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Table 4 Synergistic antifungal activity of ZnO-NPs with conventional antifungals against C. albicans and C. tropicalis. Data represent mean Inhibition zone diameters (mm ± SE, *n* = 3). Superscript letters (ᵃ, ᵇ) indicate statistical significance within columns (Tukey’s test, *p* < 0.05). IFA = Increase in fold area; ns = not significant (p ≥ 0.05). FICI (Fractional inhibitory concentration Index) interpretation: ≤0.5 synergistic, > 0.5–1.0 additive, > 1.0–4.0 indifferent, > 4.0 antagonistic.
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Fig. 13
figure 13

Proposed mechanism of synergy between ZnO NPs and nystatin: (1) membrane pore formation by nystatin, (2) enhanced uptake of ZnO NPs/Zn²⁺, (3) ROS generation, and (4) intracellular damage.

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Antioxidant activity

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 IC50 of ZnO NPs was 335.48 µg/mL whereas the IC50 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.

Fig. 14
figure 14

Concentration-dependent DPPH radical scavenging activity of biosynthesized ZnO NPs. The free radical scavenging capacity of ZnO NPs (50–1600 µg/mL) was assessed and compared to ascorbic acid as a standard antioxidant control. Data are presented as mean percentage inhibition ± standard error (SE) of three replicates.

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Cytotoxic assay of ZnONPs

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 pathways4. 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)95. 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 arrest96. 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 cells97. Recent studies also highlight the role of ZnONPs in inhibiting angiogenesis and epithelial-mesenchymal transition (EMT), thereby reducing metastasis and chemotherapy resistance in aggressive cancers98.

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 rates99. Moreover, surface functionalization with bioactive compounds (e.g., phenolic compounds from plant extracts) further modulates stability, dispersibility, and cellular interactions96. Moreover, hexagonal and spherical morphologies promote membrane disruption and internalization, while anisotropic structures like nanorods induce mechanical damage to cellular structures4. 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 pathways100.

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.

Fig. 15
figure 15

Dose-response cytotoxicity of biosynthesized ZnO NPs against HUH7 cancer and WI38 normal cell lines. Cell viability was determined by MTT assay after 48 h of treatment with various concentrations of ZnO NPs. Data are expressed as mean percentage viability ± standard error (SE) (n = 3).

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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.

Conclusions

This study successfully established a green synthesis method for zinc oxide nanoparticles (ZnO-NPs) using Lepidium sativum seed extract. The comprehensively characterized NPs demonstrated potent in vitro antifungal activity against drug-resistant C. albicans and C. tropicalis. A particularly significant finding was the strong synergistic effect observed with nystatin, suggesting a potential strategy to enhance the efficacy of conventional antifungals and overcome resistance. Furthermore, the NPs exhibited considerable antioxidant activity and selective cytotoxicity showing significantly greater potency against hepatoma (HUH7) cells (IC₅₀: 145.2 µg/mL) than against normal lung fibroblasts (WI38) (IC₅₀: 237.6 µg/mL), suggesting their potential for the development of anticancer agents mitigating chemotherapy induced disabilities. Furthermore, their intrinsic antimicrobial and anticancer properties make them compelling candidates for use as nanocarriers in targeted combination therapies.

The findings from this in vitro study pave the way for developing safer and more effective nano-based adjuvant therapies to manage recalcitrant candidiasis. However, it is important to note that these compelling findings are derived from in vitro models. The current work has limitations, including the lack of in vivo validation and assessment of efficacy against fungal biofilms, which are crucial steps toward clinical relevance. Consequently, the promising results presented here should be viewed as a foundational step that justifies further investigation.

Future research must therefore prioritize in vivo studies to confirm antifungal efficacy and systemic safety in animal models. Subsequent work should also focus on elucidating the precise molecular mechanisms of action, including ROS-mediated pathways and the basis of the observed synergy. Furthermore, future studies should explore formulating these NPs into durable coatings on polymer or metal substrates, testing their stability, biocompatibility, and efficacy in reducing biofilm formation in vivo, particularly for devices commonly used by high-risk populations, such as individuals with disabilities. Finally, based on those findings, the development of other targeted formulations, such as topical creams, should be explored. This study provides a strong rationale and a clear direction for the continued development of these multifunctional nanoparticles.