Introduction

The term “nanoflower” is used for a class of nanoscale objects characterized by a floral shape1. Nanotechnology emerged during the mid-twentieth century, whereas the idea of nanoflowers, which developed in the broader context of nanotechnology research, only rose to prominence in the late twentieth century2. The complex and eye-catching flower-like shape at the nanoscale gives these structures their name. They have distinctive geometry, consisting of petals or branches resembling natural flowers radiating outward from a central core. These structures have been synthesized from various materials, including metals and metal oxides. This unique design is achieved via a controlled synthesis procedure, including the need for seed-mediated growth, optimizing reaction conditions, and carefully selecting precursor materials3. Zinc Oxide nanoparticles (ZnO-NP) and nanoflowers (ZnO-NFs) exhibit distinctive properties at the nanoscale4. With its wide bandgap and unique characteristics, ZnO is a promising candidate for various nanomaterial applications5. Synthesis and application of nanoflowers are driving advances in fields ranging from sensors, photocatalysis, and piezoelectric devices to antibacterials6. ZnO-NFs exhibit remarkable antibacterial properties, effectively inhibiting the growth of both Gram-positive and Gram-negative bacterial strains. This attribute highlights their potential as promising agents for various antibacterial applications in healthcare and environmental settings7.

ZnO-NFs can be prepared using diverse chemical and physical methodologies, encompassing hydrothermal and solvothermal processes, which allow controlled reduction and growth. This offers morphological control through varying solvent composition and reaction conditions8. Template-assisted techniques rely on sacrificial templates to guide growth, yielding nanoflowers with distinct architectures9. Electrochemical deposition involves controlled electrodeposition onto substrates to achieve desired structures10. Microwave-assisted synthesis accelerates nucleation and growth through microwave irradiation, achieving rapid nanoflower synthesis. However, among these methods, green synthesis stands out as a superior approach since physical and chemical methods have traditionally depended on the use of hazardous chemicals and substantial energy inputs, which are environmentally undesirable. Green synthesis utilizes natural sources such as enzymes, plant extracts, microbes, or biocompatible agents for synthesis11,12,13. Utilizing biomolecules and plant extracts to synthesize nanoflowers ensures environmentally friendly procedures and introduces biocompatibility, a vital aspect for biomedical applications14,15. The biological synthesis of ZnO-NFs is complex due to its intricate hierarchical structure. Minor variations in parameters like temperature and pH lead to divergent results, hindering reproducibility. Aggregation and scaling up further complicate the process16. Thus, ongoing research and innovative approaches are essential. To the best of our knowledge, there is no known instance of ZnO-NFs synthesis involving enzymes. α-Amylase, a commercially available and efficient digestive enzyme, converts starch into simple sugars. α-Amylase has exhibited remarkable potential in our previous work on the synthesis of nanoparticles5,17. Our current research seeks to replicate this success by achieving the synthesis of ZnO-NFs. In this study, we present a novel and sustainable approach to the synthesis of ZnO-NFs using α-amylase. We employed a comprehensive experimental approach, utilizing X-ray diffraction (XRD), energy-dispersive X-ray spectroscopy (EDX), dynamic light scattering (DLS), and transmission electron microscopy (TEM) for thorough characterization. The multifaceted assessment included investigating photocatalytic, antibacterial, and antibiofilm effects of ZnO-NFs. Kinetic studies were conducted, estimating the degradation of dyes under varying pH and in the presence of interfering substances. Furthermore, we analyzed hemocompatibility against red blood cells to gauge the biocompatibility of ZnO-NFs. The present work marks a significant stride toward advancing environmentally sustainable nanomaterials, fostering a new frontier in developing ZnO-NFs with tailored properties for diverse applications.

Results and discussion

Due to its environmentally benign characteristics, biologically mediated production is a promising new approach to synthesizing novel nanomaterials. Nanoflowers, a distinctive variation of nanoparticles, offer advantages such as an enhanced surface area, unique morphology with multiple facets and edges, improved catalytic activity, potential hemocompatibility, and suitability for bioimaging3. Although enzyme-mediated synthesis of nanomaterials has been reported in the past, little study has been devoted to synthesizing nanoflowers using enzymes as biomimetic agents. Therefore, we examined the potential of α-amylase in synthesizing ZnO-NFs. When an aqueous solution of Zn(CH3COO)2·2H2O was incubated with an α-amylase enzyme solution at 25 °C and pH 9, the colorless solution turned to off-white within 4 h, indicating synthesis of ZnO-NFs. UV–Vis spectra (Fig. 1) show that the lambda max (λmax) of these particles appears at a wavelength of 360 nm, which corresponds to the surface plasmon resonance (SPR) of ZnO-NFs18.

Figure 1
figure 1

UV–Vis spectra of ZnO-NFs: (a) 0.02 M Zinc acetate + NaOH + α-amylase at pH 9, (b) 0.02 M Zinc acetate + NaOH at pH 9. Inset represents Blank (0.02 M Zinc acetate), no peak was observed.

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Since the blank [Zn(CH3COO)2·2H2O solution] did not change color over time, and no peak (inset, Fig. 1) was observed, we conclude that the enzyme is responsible for the synthesis. Parameters such as pH, enzyme concentration, and Zn(CH3COO)2·2H2O salt were also examined and optimized (Results not shown). We conducted a separate experiment in which we combined Zn(CH3COO)2·2H2O with NaOH at pH 9, we observed that the λmax exhibited a distinct difference in absorbance, characterized by a broader and less well-defined peak, compared to the peak observed with the α-amylase enzyme. The presence of a sharp and narrow peak in the UV–Vis spectrum is indicative of a highly specific and precisely defined absorption wavelength19. Because of this characteristic, we further characterized the particles synthesized by enzymatic means. DLS analysis of ZnO-NFs reveals that the size of nanoflowers in the colloidal solution is approximately 101 nm (Figure S1). The polydispersity index (PDI) was determined to assess the particle size distribution and measured at 0.186. Zeta potential analysis was employed to evaluate the stability of the colloidal solution and yielded a value of −21.2 mV ± 4.48 mV (standard deviation), indicating the stability of the colloidal solutions. In the FTIR spectrum of ZnO-NFs (Figure S2), the characteristic band at 3614 cm−1 appeared from the stretching vibration of hydroxyl (OH) groups. The characteristic band at 2351 cm−1 of C=C conjugated group. The peaks around 1635 cm−1 and 1521 cm−1 could be attributed to the strong vibration mode of Amide I and II, respectively, of the peptide backbone. Moreover, the 669 cm−1 appeared from the characteristic stretching vibration peak of the ZnO-NFs20,21.

Figure 2 illustrates an XRD pattern obtained from the synthesized ZnO nanomaterial. This pattern observed distinct diffraction peaks at specific 2θ values: 31.82, 34.41, 36.31, 47.65, 56.53, 63.07 and 67.73 degrees. These peaks can be directly attributed to the crystallographic planes denoted as (100), (002), (101), (102), (110), (103), and (112) within the hexagonal structure of ZnO-NF. This pattern closely matches the known hexagonal crystal structure (the wurtzite structure) as cataloged in the JCPDS Database under File number 36–145118.

Figure 2
figure 2

XRD pattern matching with hexagonal crystal structure (wurtzite structure) as in JCPDS Data (File number: 36-1451).

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Figure 3a shows TEM images of dried ZnO-NFs, representing an arrangement of star-shaped flowers. Each of these flower-like structures comprises a confluence of 6 to 7 wide petal-like structures. The length of the flower varies from 200–300 nm. Researchers have reported the synthesis of slightly larger nanoflowers using other processes4,22,23. Elemental analysis of ZnO-NFs was examined by EDX spectroscopy (Fig. 3b). This confirms formation of ZnO-NFs by verifying the presence of elemental Zn and O in the ZnO-NFs24. Elemental mapping revealed that the weight percentages were approximately 60.63 (zinc) and 23.93 (oxygen).

Figure 3
figure 3

(a) TEM images of dried ZnO-NFs, each of these structures comprises a confluence of 6 to 7 wide petal-like structures. (b) Representative EDX spectroscopy graph. Elemental mapping revealed the distribution of zinc and oxygen peaks.

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ZnO-NFs exhibit strong photocatalytic capabilities that render them highly effective in the decomposition of organic dyes, especially when illuminated with ultraviolet (UV) light25. To assess the photocatalytic activity of ZnO-NFs here, three commonly used dyes, methylene blue (MB), eosin (E), and reactive red (RR), were used. These dyes were exposed to a ZnO-NFs suspension with a concentration of 100 mg/L. Approximately 2 h into the experiment, a significant change in the dye's color was observed. All the dyes transitioned from vibrant to a lighter-colored solution. Maximum degradation was observed in the case of the E-dye (90%), followed by MB-dye (87%) and RR-dye (81%) after 120 min (Fig. 4). The control of each dye without ZnO-NFs under UV light exhibited negligible degradation. By varying the concentration of ZnO-NFs, we observed that an increase in ZnO-NFs load led to a decrease in degradation efficiency. This decline is likely due to a reduction in available reactive surface sites, potentially impeding light penetration and reducing the number of photons reaching the active sites of the photocatalyst26. The photocatalytic reaction typically encompasses photoexcitation, charge separation, migration, and surface oxidation–reduction reactions27. During illumination, the predominant reactive species generated are h+ (positive holes), •O2− (superoxide radicals), and OH− (hydroxide ions). It begins with photoelectrons (e−) transitioning from the valence band to the conduction band, leaving behind positive holes (h+) in the valence band. These holes subsequently interact with water molecules, instigating the formation of hydroxyl free radicals. These hydroxyl radicals play a pivotal role in the degradation process of dye molecules28,29.

Figure 4
figure 4

Dye degradation efficiency of ZnO-NFs versus time against methylene blue, eosin, and reactive red. Maximum degradation was observed in the case of eosin (90% approx.). Values are means of three replicates ± standard deviation.

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The Langmuir–Hinshelwood kinetic model was utilized to calculate the rate of photocatalytic degradation with respect to dye concentration30,31. The degradation kinetics of MB, E, and RR followed a pseudo-first-order model. The resulting rate constants and R2 values derived from the dye degradation studies are presented in Supplementary data (Table 1).

The effect of pH on dye degradation was minimal up to pH 10. However, beyond pH 10, there was a slight decline (~ 15 and ~ 10%) in the percentage degradation for MB and E dye, respectively, while no significant change was observed in the degradation of RR dye (Figure S3). The pH of the solution plays a crucial role in photocatalytic reactions occurring on particle surfaces, influencing both the surface charge and the formation of aggregates. The degradation was monitored at the λmax for each dye. Figure S3 also reveals a correlation between the concentration of various interfering substances and the percentage degradation of the dye. NO3, Fe2+, and humic acid exhibited insignificant effects on dye degradation. Contrastingly, CI− resulted in approximately 5% and 15% reduction in the degradation percentages of RR and MB dye, respectively, when subjected to ZnO-NFs. Ca2+ demonstrated an approximately 10% reduction in the degradation percentages of MB and E dye, along with a roughly 15% reduction in the degradation percentage of RR dye. The inhibitory effects of Cl− and Ca2+ ions on dye degradation suggest a competitive interaction with dye molecules for photo-oxidizing species on the catalyst's surface, hindering dye degradation by ROS32,33.

ZnO-NFs were also analyzed for their antibacterial properties against Staphylococcus aureus, Escherichia coli, Pseudomonas aeruginosa, Streptococcus mutans, Enterococcus faecalis, and Klebsiella pneumoniae. Additionally, its antibiofilm properties were investigated against select strains from this group. These bacterial species are commonly associated with various types of infections. Staphylococcus aureus can cause skin infections and bloodstream infections. Escherichia coli and Enterococcus faecalis are known for causing urinary tract infections. Pseudomonas aeruginosa is often associated with lung infections. Streptococcus mutans is a major contributor to dental cavities and tooth decay. Klebsiella pneumoniae commonly causes pneumonia34,35,36. The S. aureus used in this study was a penicillin-resistant strain. These species can become resistant to antibiotics over time, making infections more difficult to treat in clinical settings. As previously mentioned, ZnO-NFs produce reactive oxygen species (ROS) when exposed to light. These ROS have the potential to kill bacterial cells, even those that are resistant to antibiotics37. ZnO-NFs can also release Zn2⁺ ions, which have the ability to disrupt extracellular polymeric substances (EPS) that form the backbone of biofilms7. Characteristics of biofilms, such as their resistance to antibiotic penetration and influence on microbial physiology, make it difficult for conventional antibiotics to successfully eradicate bacteria in these environments38. As a result, this encourages persistent infections which may become progressively less responsive to treatment. Additionally, exchange of genetic material within biofilms facilitates transmission of resistance genes, contributing to multidrug-resistance39. Figure 5a; depicts inhibition of cell viability expressed as a percentage for the three bacterial strains following a 24-h treatment with ZnO-NFs. At a concentration of 100 mg/L, ZnO-NFs demonstrated significant viability inhibition, with the highest effect observed in the case of S. aureus (92%), E. coli (82%), P. aeruginosa (67%), S. mutans (90%), E. faecalis (85%) and K. pneumonia (78%). The surface area of ZnO-NFs also influences the biocidal effect. The petal-like structure is known to have a larger surface area, offering more functional sites for antibacterial behavior40. The well-defined, pointed edges present on ZnO-NFs serve as physical puncturing agents, causing damage to bacterial cell walls and membranes. This mechanical disruption upsets bacterial structural integrity, leading to cell lysis41. In order to assess bacterial cell density, we conducted SEM analysis on both the 0-h and 24-h E. coli cultures treated with 100 mg/L ZnO-NFs. Figure 5b and S4 illustrate SEM images of the E. coli and E. faecalis cultures, respectively, subjected to ZnO-NFs treatment, clearly depicting a significant reduction in bacterial cell density at the 24-h time point. This observation shows that ZnO-NFs exhibits potent antibacterial properties.

Figure 5
figure 5

(a) Cell viability inhibition assay of ZnO-NFs against E. coli, S. aureus, and P. aeruginosa. Values are means of three replicates ± Standard Deviation, ANOVA significant at p ≤ 0.05. (b) SEM images of the E. coli cultures subjected to ZnO-NFs treatment showed a significant reduction in bacterial cell density at a 24-h time point, compared to 0-h.

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Figure S5a represents the biofilm inhibition capacity of ZnO-NFs at different concentrations against biofilms of S. aureus, E. coli, and P. aeruginosa. At 200 mg/L of ZnO-NFs, maximum biofilm inhibition was observed in the case of P. aeruginosa (89.2%) followed by E. coli (85.6%) and S. aureus (66%). The variation in the percentage inhibition of biofilms in the present study may be attributable to differences in composition and maturity of the biofilms. These factors can significantly influence biofilms' resistance levels, as mature and densely structured biofilms typically present greater challenges in terms of elimination42. Figure S5b represents the SEM image of biofilm of S. aureus, it can be observed that the formation of biofilm has been inhibited by the treatment of ZnO-NFs.

Ability of ZnO-NFs to generate ROS has been established and further confirmed by a nitroblue tetrazolium (NBT) assay. Figure 6 shows a significant increase in ROS production by ZnO-NFs under UV light, contrasting with notably lower ROS production observed under dark conditions.

Figure 6
figure 6

ROS activity was measured using NBT after 5 h of exposure to UV light and in the dark. Data shown are mean ± standard deviation. All experiments were performed in triplicates; mean ± SD are shown, having the p-value < 0.05.

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In Table 1, a comparative presentation highlights the correlation between the current study's results and prior research on ZnO-NFs synthesis and application. Figure 7 provides an overview of the capacity of ZnO-NFs to damage the membranes of red blood cells, which was evaluated by measuring the release of haemoglobin into the medium in a hemolysis assay. When ZnO-NFs were introduced at a concentration of 600 ppm, the resulting hemolysis is only 1.79%, indicating a non-hemolytic response. However, at 800 ppm, the hemolysis rate increased to 3.96%, suggesting a slightly hemolytic effect. These results align with the American Society for Testing and Materials (ASTM) F756-00 standard, which designates a hemolysis (H) degree as (H = 0–2%, non-hemolytic; H = 2–5%, slightly hemolytic; H ≥ 5%, hemolytic)43,44. In contrast, Triton-X (used as a positive control) leads to a significant hemolysis rate of 98%. Consequently, it can be concluded that ZnO-NFs, up to a concentration of 600 ppm, exhibit significant hemocompatibility. This is an important factor for materials that may eventually encounter blood in potential future applications in biomedical devices or drug delivery systems45.

Figure 7
figure 7

Hemolysis Rate of ZnO-NFs. The rate of hemolysis remains lower than 2% (1.79%) until 600 ppm for 24 h. Triton-X (positive control) leads to a significant hemolysis rate of 98%. The results are the mean ± SD of triplicate experiments. ANOVA significant at p ≤ 0.05.

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Conclusion

In this study, we explored the enzymatic synthesis of ZnO-NFs using α-amylase, emphasizing their potential. Structural characterization via X-ray diffraction confirmed a hexagonal crystal structure, while TEM images vividly illustrated their star-shaped morphology. Our findings highlight the multifaceted potential of ZnO-NFs, including photocatalytic efficiency, antibacterial properties, and biocompatibility. ZnO-NFs exhibited significant efficiency in degrading organic dyes and displayed remarkable antibacterial and antibiofilm properties, including against a penicillin-resistant S. aureus strain. Moreover, their inhibition of biofilms, particularly in P. aeruginosa, underscores their potential against antibiotic-resistant strains. ZnO-NFs also demonstrated remarkable hemocompatibility up to a concentration of 600 ppm. The enzyme-mediated synthesis of nanoflowers opens avenues for sustainable and controlled materials synthesis, offering promising prospects for future innovations in healthcare and nanotechnology. Further optimization of the enzymatic synthesis process could enhance scalability and facilitate the synthesis of nanoflowers with other metal salts. Additionally, investigating the long-term stability and performance of ZnO-NFs in real-world applications could contribute to the ongoing development of sustainable nanomaterials for practical use.

Materials and methods

Chemicals

Zinc acetate dihydrate (Zn(CH3COO)2.2H2O, > %98), sodium hydroxide (NaOH, > %99), triton X-100, ascorbic acid, culture media were obtained from Sigma Aldrich, Life Science, USA. Distilled water (DI) was produced using a two-stage Millipore Direct-Q3. α-amylase was purchased from Hi-media (India). Escherichia coli (ATCC-8739), Staphylococcus aureus (ATCC 11,632), Streptococcus mutans (ATCC 25,175), Pseudomonas aeruginosa (ATCC 27,853) were obtained from the American Type Culture Collection, USA, whilst Enterococcus faecalis (MTCC 439) and Klebsiella pneumoniae (MTCC 109) strains were procured from the Microbial Type Culture Collection (MTCC), Institute of Microbial Technology (Chandigarh, India). All other chemicals and reagents were purchased from Sigma Aldrich, Life Science USA and were of analytical grade with no further purification required.

Synthesis of ZnO-NFs

ZnO-NFs were synthesized by incubating 30 mL of α-amylase (1 mg/mL in distilled water) with 70 mL of a freshly prepared aqueous solution of 0.02 M Zinc acetate dihydrate. This mixing was carried out in a dropwise manner at 25 °C and at pH 9. Appropriate aliquots were withdrawn after regular time intervals (30 min) and synthesis was checked using UV–VIS spectroscopy. After 4 h, the reaction mixture was centrifuged at 3000 × g for 10 min, and the pellet containing ZnO-NFs was washed with distilled water, followed by drying. After drying, the nanoparticles were used for further characterization. Parameters such as concentration of enzyme and pH were also optimized. A control was run in which only 0.02 M Zinc acetate dihydrate solution was taken without α-amylase and the spectra were recorded after regular intervals of time.

Characterization of ZnO-NFs

UV–Vis spectroscopy

ZnO-NFs were initially characterized using UV–visible spectroscopy (Mecasys Optizen 3220 UV spectrophotometer). An aliquot of 1 mL was removed from the solution during the synthesis of ZnO-NFs, and spectroscopic measurements were taken in the wavelength range of 300–700 nm.

Dynamic light scattering (DLS) and Fourier-transform infrared (FTIR) spectroscopy

A Malvern Zetasizer Nano ZS was utilized for DLS measurements. The analysis was carried out at a consistent temperature of 25 °C throughout 10 cycles. ZnO-NFs at a concentration of 1 mg/ml, dissolved in distilled water, were used for the analysis. For FTIR, ZnO-NFs in the form of powder were used. FTIR spectra were recorded with a Shimadzu, FTIR between 4000 and 550 cm−1, with a resolution of 4 cm−1.

X-ray diffraction (XRD)

XRD analysis was performed on a Bruker D8 advance diffractometer over a wide range of Bragg angles (20° ≤ 2θ ≤ 80°) as described previously5.

Transmission electron microscopy (TEM)

TEM analysis of dried ZnO-NFs was performed on a JEOL, F2100 instrument as described earlier5. For elemental analysis, Energy dispersive X-ray spectroscopy (EDX) (Model EVO-40, ZEISS) was used.

Photocatalytic activity of ZnO-NFs under UV light

The degradation efficiency of three different dyes, MB, E, and RR, was assessed to evaluate the photocatalytic performance of ZnO-NFs under ultraviolet (UV) light irradiation as described previously with slight modification46. A total of 70 mL of each dye solution (25 mg/L) was mixed with 30 mL of ZnO-NFs suspension, (100 mg/L). The mixture was subjected to sonication and subsequently placed inside a UV incubator. A control experiment was conducted under the same conditions, using solutions of each dye at the same concentration without the presence of ZnO-NFs. The absorbance of the solutions was monitored at various time intervals for a duration of up to 2 h. The experiment was run in triplicate. The degradation percentage of the dyes was determined using the below equation.

$${\text{Degradation}}\;(\% ) = \left[ {\left( {{\text{C}}_{{\text{o}}} - {\text{C}}_{{\text{t}}} } \right)/{\text{C}}_{{\text{o}}} } \right] \times 100,$$

where Ct is the absorbance at time t, and Co is the initial absorbance at time t0.

Interference study

The effects of pH (3–13), humic acid (25 ppm), various interfering ions Cl− (0–1000 ppm), NO3− (0–1000 ppm), Ca2+ (0–200 ppm) and Fe2+ (0–200 ppm) on dye degradation was analyzed. The pH was adjusted by adding 1N HNO3 or 1N NaOH. 30 ppm of each dye was used to interact with ZnO-NFs, in the pH range of 3–13 under UV irradiation for 60 min.

Bacterial cell viability

Five mL of freshly cultured bacterial cells (S. aureus, E. coli, P. aeruginosa, S. mutans, E. faecalis and K. pneumoniae) with an initial concentration of 106 CFU/mL were incubated as described previously47 in an appropriate nutrient-rich medium containing ZnO-NFs at concentrations ranging from 20 to 100 mg/L. The cultures were maintained at a temperature of 37 °C for a continuous period of 24 h with constant agitation. After 24 h, optical density was measured at 600 nm by taking aliquots (1 mL) from the culture, using a spectrophotometer. These experiments were carried out in triplicate. Additionally, control groups for each bacterial culture, maintained under the same conditions but without the presence of ZnO-NFs, were included. Optical density measurements for these control cultures were obtained after the same 24 h incubation period. Cell viability was determined by calculating the percentage of viable cells in the control sample, which was considered 100%.

Cell density studies

The cell density of E. coli and E. faecalis bacterial strains was assessed both prior to and subsequent to the treatment with ZnO-NFs. Samples from the bacterial culture were taken and fixed onto a glass slide using 2.5% glutaraldehyde and 2% paraformaldehyde (prepared in a 0.1 M sodium phosphate buffer, pH 7.4 at 4 °C). Subsequently, cells were washed three times with phosphate buffered saline (PBS) solution. For SEM analysis, a section of the glass slide was subjected to gold coating through cathode spraying and subsequently examined utilizing a SEM (Leo 435 VP model equipped with digital imaging and a 35 mm photography system).

Biofilm inhibition

The capacity of ZnO–NFs to inhibit the growth of biofilms was assessed using E. coli, S. aureus, and P. aeruginosa as the test microorganisms by crystal violet (CV) assay48. Inoculum was prepared from overnight-grown cultures, diluted to OD600 = 0.01. Thereafter, 1 mL of each of the bacterial cultures and 1 mL of culture medium were inoculated in each well of a 24-well plate, and the plate was incubated at 37 °C for 24 h. Various concentrations of ZnO-NFs (50, 100, 150, 200, 250 mg/L) were added and incubated with the test microorganisms and kept at 37 °C for 48 h. After incubation, biofilms were rinsed three times with distilled water to remove any non-adhered cells. Crystal violet (CV) was introduced into the wells and allowed to incubate for 60 min. Subsequently, CV was removed by washing it with water. Bacterial cultures without the ZnO-NFs treatment were considered as controls. The amount of total biomass was determined by measuring the absorbance at 595 nm using a Multiskan™ FC Microplate reader. The experiment was performed in triplicate. Wells with only the bacterial culture served as negative controls. The percentage inhibition of biofilms was measured using the equation below.

$${\text{Inhibition}}\;{\text{of}}\;{\text{biofilm}}\;(\% ) = \left[ {\left( {{\text{C}} - {\text{B}}} \right){-}\left( {{\text{T}} - {\text{B}}} \right)/\left( {{\text{C}} - {\text{B}}} \right)} \right] \times 100,$$

where B = average absorbance of every well for blank (empty wells), C = average absorbance of every well for control wells (bacterial culture), T = average absorbance of every well for treated wells (bacterial culture + ZnO-NFs).

Production of ROS

The quantification of ROS generated by ZnO-NFs was carried out by NBT Assay49, a colorimetric method specifically designed for the assessment of ROS production. This assay involves the conversion of water-soluble NBT salt into the water-insoluble, blue NBT-diformazan in the presence of ROS. ZnO-NFs at a concentration of 1 mg/ml were combined with NBT in a volumetric ratio of 20 parts ZnO-NFs suspension to 2 parts NBT solution. The resultant samples were incubated for 5 h, under both UV light and dark conditions. Subsequently, absorbance measurements were taken at 450 nm.

Hemolytic assays

Hemolysis assay of ZnO-NFs was performed as previously described with slight modifications50. 5 mL of fresh human blood was obtained from a healthy male donor (25 years). Erythrocytes were isolated by centrifugation at 1,200 g for 10 min, washed three times in PBS buffer, and then diluted to a final concentration of 5%. A 100μL aliquot of erythrocytes was placed in a 96-well cell culture plate together with ZnO-NFs (100–1000 ppm). All measurements were performed in triplicate. The plate was incubated for 1 h at 37 °C, gently agitating it at 150 rpm. The plate was then centrifuged for 10 min at 1,500 g. A fresh 96-well plate was used to transfer aliquots (50 μL) of supernatant, which were then diluted with an additional 50 μL of PBS buffer. Hemolytic activity was determined by absorbance measurement at 540 nm with a Multiskan™ FC Microplate reader. We employed Triton X-100 (0.1% in PBS), which has the capacity to fully break down red blood cells, as the positive control, while we used PBS as the negative control. The hemolysis percentage was calculated using the following equation:

$${\text{Hemolysis}}\;(\% ) = \frac{{{\text{OD}}_{{{\text{SAMPLE}}}} - {\text{OD}}_{{{\text{NEGATIVE}}}} }}{{{\text{OD}}_{{{\text{POSITIVE}}}} - {\text{OD}}_{{{\text{NEGATIVE}}}} }}$$

Statistical analysis

The data shown are the mean of three replicates (n = 3) ± standard deviation. Data entry was performed in a Microsoft EXCEL spreadsheet and final analysis was performed using Statistical Package for Social Sciences (SPSS) software, IBM manufacturer, Chicago, USA, ver 25.0. One-way ANOVA or independent-sample t-tests were conducted to analyze significant differences. In the case of the rest of the data, statistical significance was taken as p < 0.05.

Table 1 Comparative analysis of current study findings with previously published literature on ZnO-NFs.
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