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. 2026 Mar 9;16(4):122. doi: 10.1007/s13205-026-04754-7

Thin films coating with ZnO nanoparticles synthesized via acetic acid bacteria: Structural and antibacterial characterization for food applications

Gülden Kılıç 1,✉
PMCID: PMC12968136  PMID: 41808724

Abstract

In this study, zinc oxide nanoparticles (ZnO NPs) were synthesized using extracellular metabolites of acetic acid bacteria (AAB) derived from fermented beverages, and then a thin film coating of ZnO NPs was created via spin coating techniques. ZnO NPs and thin films obtained were characterized using physical, structural, and biological techniques. Firstly, two AAB isolates were obtained from olive leaf vinegar and kombucha with ginger, and genotypically identified. ZnO NPs synthesis by N. hansenii B1 and N. hansenii B2 was indicated by a white precipitate and confirmed for both NPs and thin films by UV-Vis absorption at 300–400 nm. SEM images revealed agglomerated ZnO NPs and homogeneous nanosized thin films, while EDX analysis confirmed their pure phase composition with zinc and oxygen peaks. XRD results showed the presence of monoclinic and cubic crystal ZnO NPs, with crystallite sizes ranging from 9.68 to 35.11 nm, while the thin films were amorphous. FTIR spectra revealed 519 and 3350 cm− 1 peaks corresponding to ZnO bonds. The measurements of water contact angles of ZnO thin films exhibited hydrophilic surface characteristics. Moreover, ZnO NPs exhibited strong antibacterial activity, being effective against S. aureus and E. coli O157:H7 (12.50–14.00 mm), with minimum inhibitory concentration values of 0.625–1.25 mg/mL. The inhibition zones of thin films were found between 0.50 and 2.38 mm, and both films completely inhibited E. coli O157:H7 in direct contact tests after 2 h. These findings suggest that green-synthesized ZnO NPs and their thin films have great potential as antimicrobial applications in various food, medicine, and related fields.

Supplementary Information

The online version contains supplementary material available at 10.1007/s13205-026-04754-7.

Keywords: Microbial synthesis, Fermented foods, Microbiota, Spin coating, Antimicrobial

Introduction

Nanotechnology, which involves the manipulation of atomic, molecular, and macromolecular matter to the nanometer scale, has emerged as a highly promising field of study in the twenty-first century, with diverse applications in biotechnology, chemistry, medicine, and material sciences (Ahmed et al. 2024). However, nanotechnology has now become deeply integrated into real-life applications across multiple sectors. In medicine, nanoparticles are used for targeted drug delivery, cancer imaging, antibacterial wound dressings, and controlled-release therapeutic systems (Kianfar 2021; Hlaváček et al. 2022; Gültekin et al. 2024; Lajmiri et al. 2024). In the food industry, nanostructures contribute to active and intelligent packaging, nanosensors for real-time spoilage detection, antimicrobial coatings for food-contact surfaces, and nutrient fortification (Elshayb et al. 2021; Sharma et al. 2023; Jayakumar et al. 2023; Gökmen et al. 2024). Environmental applications include photocatalytic water purification, pollutant degradation, and membrane-based filtration technologies (Shahid-ul-Islam et al. 2023; Geldasa et al. 2023; Shukla et al. 2024). Consumer products such as sunscreen, textiles, electronics, and cosmetics also employ nanomaterials to improve UV protection, durability, and functional performance (Xiao et al. 2022; Rabiei et al. 2022; Sharma et al. 2023). NPs, described as particles with at least one dimension in 1–100 nm, show special physicochemical properties that vary significantly with the size. Various metal or metal oxide-based NPs, such as aluminum (Al), gold (Au), iron (Fe), iron oxide (FeO), selenium (Se), silver (Ag), zinc (Zn), and zinc oxide (ZnO), have been synthesized through both bottom-up and top-down approaches (Singh et al. 2021; Kirubakaran et al. 2025). The bottom-up approach assembles smaller molecular components into NPs, while the top-down approach involves techniques such as electro-explosion, etching, laser ablation, lithographic processes, and milling (Fu et al. 2018). These methods allow for precise control over NP stability and morphology, making them appropriate for different applications (Jamkhande et al. 2019). However, these synthesis methods require expensive equipment and involve dangerous chemicals, raising concerns about toxicity to humans and the environment and limiting large-scale production (Siaw et al. 2020; Kirubakaran et al. 2025). Hence, the green synthesis of NPs has attracted great attention as a sustainable approach because of its biocompatibility, cost-effectiveness, low toxicity, and minimal environmental impact in recent years. This current approach for NP synthesis is characterized by the absence of dangerous chemicals, high temperatures, and pressure, providing both human health and environmental safety (Mariotti et al. 2020). This approach uses naturally occurring biological materials, including microorganisms (bacteria, mold, yeast, virus, or microalgae) and plant-based materials (leaf root, fruit, seed and/or stem, etc.) or molecules (organic acids, alkaloids, flavonoids, polysaccharides, proteins, vitamins, terpenoids, and certain heterocyclic compounds, etc.) to facilitate NP synthesis in an economically and ecologically viable manner (Gahlawat and Choudhury 2019; Rana et al. 2020). In this method, microorganisms or extracts generally react with the metallic salt, followed by the biological reduction, which is performed to convert the metal to NPs. The obtained NPs are suitable for utilization after being characterized via various techniques (Mittal et al. 2013; Ali et al. 2019). Microorganisms have been widely studied for their ability to biosynthesize various NPs through biochemical mechanisms (Pandit et al. 2022). One of the key pathways for NPs biosynthesis under dark conditions involves the NADH-dependent enzymatic bioreduction of metal ions into nanometals or nanometal oxides (Mohd Yusof et al. 2019). This biosynthesis can occur both intracellularly and extracellularly. In extracellular synthesis, the culture filtrate is obtained by centrifugation and mixed with a metallic salt solution, with NP formation indicated by a color change, such as from light yellow to dark brown for Ag NPs (Hossain et al. 2019; Ibrahim et al. 2020). In intracellular synthesis, microbial biomass is thoroughly washed and incubated with a metal ion solution under optimal growth conditions. The formation of NPs is also signaled by a color change, after which NPs are collected via sonication, centrifugation, and washing (Soni et al. 2018). However, since the intracellular method requires additional steps, the extracellular method is more advantageous (Mahdi et al. 2021).

Among microbial sources for NP synthesis, beneficial microorganisms are frequently utilized in food fermentations and nutraceutical applications. These microorganisms, known for their positive effects on health, include lactic acid bacteria (LAB), yeast, and AAB. These microorganisms have been used as starter cultures for various fermented foods, such as pickles, vinegar, wine, beer, kombucha, shalgam, and kefir etc. (Hata et al. 2023; De Bellis and Rizzello 2024). These properties make them very interesting biological sources for NP synthesis and delivery. In many studies, Lactobacillus sp., L. acidophilus, Glucanoacetobacter kombuchae, species commonly found in fermented foods, have already been employed in the biosynthesis of NPs, including Ag, Fe2O3, and Se NPs (Faramarzi et al. 2020; Majumder et al. 2022; Mohammed et al. 2023; Abd Qasim and Yaaqoob 2023). These biogenic NPs derived from beneficial microorganisms have shown potential in various fields, such as antimicrobial and anticancer agents, in agriculture, drug delivery systems, and the food industry (Gomez-Zavaglia et al. 2022; Shanmugam et al. 2023).

ZnO NPs have garnered significant interest due to their remarkable antimicrobial properties, UV shielding capabilities, and high surface area, making them valuable for applications in diverse fields, including biomedicine, food preservation, and environmental remediation (Gökmen et al. 2024). On the other hand, although ZnO is best known for its thermodynamically stable hexagonal wurtzite structure, several studies have shown that ZnO can also crystallize in metastable cubic phases (most commonly the zinc blende (sphalerite) and, less frequently, the rock-salt structure) under specific physicochemical conditions (Zarhri et al. 2022; Jobe et al. 2022; Subramani et al. 2023). The formation of cubic ZnO has been associated with nanoscale confinement, low-temperature synthesis, biomolecule-mediated nucleation, and biological or plant extract-assisted green synthesis routes, where organic moieties can alter growth kinetics and selectively stabilize non-wurtzite lattices (Francis 2024; Saw et al. 2024). Cubic ZnO nanostructures exhibit distinct surface energies, defect distributions, and band-gap characteristics relative to the wurtzite phase, which can influence their optical absorption behavior, excitonic transitions, catalytic activity, and antimicrobial properties. The stabilization of such metastable cubic polymorphs therefore provides an opportunity to tailor functional performance and has been increasingly reported in green-synthesized and solution-processed nanomaterials (Abel et al. 2021; Meky et al. 2023; Naseer and Iqbal 2024; Bhandari et al. 2025). Also, ZnO NPs are currently described as a “generally recognised as safe (GRAS)” material by the U.S. Food and Drug Administration (FDA) and can also be used as a food additive. ZnO NPs are reported by several scientists to be harmless (non-toxic) to human cells (Shi et al. 2014; Kadhim et al. 2019). Moreover, it has been stated that with the increasing global challenge of foodborne pathogens and antimicrobial resistance, exploring biogenic ZnO NPs as safe and effective antimicrobial agents is highly relevant (Gökmen et al. 2024). In this regard, the synthesis and characterization of biogenic ZnO NPs from reliable and safe sources is of significant relevance. Recently, in different studies, ZnO NPs were synthesized as green synthesis via microorganisms (Bacillus sp., B. subtilis, Weissella cibria, W. confusa, Lactobacillus gasseri, Lactococcus lactis, Pediococcus sp., Saccharomyces cerevisiae) and characterized with biological properties such as anticancer, antimicrobial, and anti-inflammatory (Mahdi et al. 2021; El-Sayed et al. 2021; Todorov et al. 2022; Al-Tameemi et al. 2023; Hamk et al. 2023; Al-Tameemi et al. 2023; El-Khawaga et al. 2025; AL-Tameemi et al. 2025).

Incorporating metal or metal oxide nanoparticles into food packaging enhances mechanical and optical properties, including UV protection, while also inhibiting microbial growth and biofilm formation on food-contact surfaces (Lomate et al. 2018; Torres Dominguez et al. 2020). Glass is widely used in equipment for the medical and food industries due to its smooth, non-porous surface, which makes it easy to clean and limits bacterial growth. Its transparency allows for visual inspection, and its chemical inertness and high thermal stability make it ideal for food processing and sterilization. Additionally, glass is recyclable, supporting sustainable practices. Recent studies indicated that the surface topography and nanostructures of glass can influence bacterial adhesion and metabolism, affect bioactivity, and highlight its potential for antimicrobial applications (Orava et al. 2014). Bioactive glasses (BAGs), which are mainly composed of SiO2, Na2O, CaO, and P2O5, have demonstrated antibacterial properties and are being investigated for use in orthopedic and dental implants (Drago et al. 2018). On the other hand, ZnO prepared via the sol-gel method is commonly used in spin or dip coating for thin film fabrication due to its low cost and simplicity. It allows for the formation of uniform nanostructures even at low temperatures (Benrezgua et al. 2022). Transition metal doping, particularly with Cu, has been extensively studied to enhance properties such as transparency, ferromagnetism, crystallinity, and conductivity (Al-Khanbashi et al. 2014; Ali et al. 2019; Dabir et al. 2020). Consequently, integrating ZnO NPs into antimicrobial coating is seen as a promising strategy for mitigating microbial contamination while ensuring biocompatibility.

Although microbial routes for ZnO NP synthesis have been reported, existing studies largely focus on plant extracts or a limited range of bacterial and fungal species, with little attention given to AAB, particularly those isolated from traditional fermented beverages. To the best of our knowledge, the use of AAB derived from vinegar, kombucha, and beetroot kvass as biogenic agents for ZnO NP synthesis, followed by their integration into functional thin films, has not been systematically investigated. This study addresses this gap by introducing a sustainable and food-origin microbial platform for ZnO NP production and demonstrating its applicability in thin film fabrication via spin coating. Specifically, this study aims to (i) synthesize ZnO NPs using AAB isolated from fermented foods, ii) create a thin film coating of ZnO NPs using spin coating techniques iii) characterize NPs and thin films through Ultraviolet-Visible (UV-Vis) spectroscopy, Scanning Electron Microscopy (SEM), Energy-Dispersive analysis X-Ray (EDX), X-Ray Diffraction (XRD), Fourier Transform Infrared (FTIR) spectroscopy, and goniometer and iv) investigate the antimicrobial efficacy of ZnO NPs and thin films produced against Staphylococcus aureus 6538P and Escherichia coli O157:H7 ATCC 35150 to determine their potential as novel antimicrobial agents. The combined use of fermented beverage-derived AAB and thin film fabrication offers a novel and scalable approach for developing antimicrobial ZnO-based coatings.

Materials and methods

Collection of samples

In this study, various fermented beverages such as vinegar [ginger vinegar (S1) and olive leaf vinegar (S2)], kombucha [produced with ginger (S3) and black mulberry (S4)], and beetroot kvass [produced with red beetroot (S5) and red beet+ginger (S6)] beverages produced by spontaneous fermentation were used (Table 1), and the samples were supplied from local producers in Alanya, Antalya (Türkiye). At each sampling point, samples were collected in original bottles and transported to a laboratory at 4 °C within 24 h. Also, the schematic representation of the experimantal workflow was given in Fig. 1.

Table 1.

Fermented beverages and their microbiological properties

Samples Sample code Ingredients AAB counts (log CFU/mL)
Ginger vinegar S1 Fresh ginger, water 2.88 ± 0.03a
Olive leaf vinegar S2 Wild olive leaves, water 3.31 ± 0.01b
Kombucha with ginger S3 Green tea, fresh ginger, sugar beet, water 5.22 ± 0.01e
Kombucha with black mulberry S4 Green tea, blackberry juice, sugar beet, water 5.10 ± 0.05d
Beetroot kvass S5 Red beetroot, rock salt, kefir whey, water 4.95 ± 0.05c
Beetroot kvass with ginger S6 Red beetroot, rock salt, fresh ginger, kefir whey, water 5.05 ± 0.03d
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*Means in the same column with different lowercase (a, b, c, d) are significantly different (P < 0.05)

Fig. 1.

Fig. 1

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Schematic representation of the experimental workflow

Microbiological properties of beverages: Isolation and identification

Decimal dilutions of the beverages were prepared, followed by plating on growth media in duplicates for microbial counts. AAB counts were determined by spread plate method on Glucose Yeast Extract Calcium Carbonate (GYC) agar (pH 6.8 ± 0.2), containing 10% D-glucose (Biolife), 1% yeast extract (Oxoid), 2% CaCO3 (Tekkim), 3% ethanol (Sigma-Aldrich), 0.5% glacial acetic acid (Merck), 1.5% agar (Biolife), and the plates were incubated at 30 °C for 5–10 days (Vero et al. 2013).

After incubation, colonies with a cream colour and a clean precipitation zone on GYC agar were considered typical AABs. Two AAB colonies were randomly selected from the plates and checked for purity by streaking them onto GYC agar. For long-term storage, the strains were kept at -18 °C in GYC broth containing 50% (v/v) glycerol.

The 16S rRNA gene sequences of two AAB isolates were amplified using universal primers 27F (5′-AGA GTT TGA TCA TGG CTC AG-3′) as a forward primer and 1492R (5′-GGT TAC CTT GTT ACG ACT T-3′) as a reverse primer. The High Pure PCR Template Preparation Kit (Sigma-Aldrich) was used for DNA extraction from AAB isolates. The PCR analysis of 16S rRNA was performed using the Xpert Fast Hotstart 2X Mastermix Kit (Grisp) with appropriate universal primers (27F-1492R).

The DNA sequencing was conducted using Sanger sequencing, and the sequences of AAB isolates were compared with reference strains using the Basic Local Alignment Search Tool (BLAST) programme at the National Centre for Biotechnology Information (NCBI) and identified. Finally, the sequences were submitted to GenBank (https://submit.ncbi.nlm.nih.gov) at the NCBI (Benson et al. 2013) under the accession numbers as PV200304 (B1 strain), and PV200774 (B2 strain).

The evolutionary analysis was performed in MEGA X (Kumar et al. 2018). The evolutionary history was inferred using the Neighbour-Joining Method (Saitou and Nei 1987). The optimal phylogenetic trees for AAB isolates were constructed separately (Fig. 2). Evolutionary distances were calculated using the Tamura-Nei method and expressed in terms of the number of base substitutions per site (Felsenstein 1985; Tamura and Nei 1993). This analysis included 11 nucleotide sequences. In this context, for the phylogenetic tree of two strains, the sequence of the outgroup (Salmonella Typhimurium ATCC 13311) was obtained from the NCBI database, the type strains (Novacetimonas hansenii strain Gachhui RG3, Novacetimonas hansenii strain LMG 1527, Novacetimonas hansenii strain NBRC 14820, Komagataeibacter saccharivorans strain LMG 1582, Komagataeibacter saccharivorans strain LMG 18909, Komagataeibacter saccharivorans strain DSM 16373, Gluconacetobacter diazotrophicus strain LMG 7603, Acetobacter syzygii strain NBRC 16604) were also retrieved from the NCBI database and included in the trees to show the relatedness of AAB isolates with the other AAB strains.

Fig. 2.

Fig. 2

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Phylogenetic tree showing the relationships between AAB strains isolated from fermented beverages and reference type strains. The tree was constructed based on 16 S rRNA gene sequence analysis, illustrating the taxonomic positions of B1 and B2 isolates used for ZnO NPs synthesis

Microbial synthesis of ZnO NPs

The synthesis of ZnO NPs was accomplished using two AAB strains identified as described above. The cultures were activated in GYC broth at 30 °C for 5–10 days. The cultures were then centrifuged at 10,000 rpm for 15 min to separate the cell supernatant. The cell-free supernatants were used to synthesize ZnO NPs (extracellular synthesis), and the NP synthesis was performed according to the study of Ramesh et al. (2021), with some modifications.

The culture supernatants (100 mL) adjusted to pH 7.5 were separately mixed with 50 mL of zinc acetate (0.1 M) used as a precursor in a 250 mL Erlenmeyer flask. The resulting solutions were incubated at 37 °C for 24 h in a shaking incubator at 150 rpm. After the color of the solutions changed from light brown to pale and deep white (the color change indicates NP synthesis), the reaction mixture was centrifuged at 10,000 rpm for 15 min at 4 °C to remove the supernatant. The supernatant was then replaced with distilled water, and the centrifugation process was repeated three more times under the same conditions to remove any remaining supernatant. The pellet-shaped collection of NPs (or NP biomass) was then dried in an oven at 40 °C for 18–24 h. The dry powder was carefully gathered and stored for the following analysis (Ramesh et al. 2021).

The coating process of ZnO NPs

The spin coating process was conducted to coat the glass substrates. Laurell WS-400BZ-6NPP/LITE (REV. MS) model spin coater was used to prepare thin films. To prepare the glass substrates, microscope glasses were cut into 2 cm × 2 cm sizes and washed by using an ultrasonicator for 15 min with acetone (Merck, Germany), isopropanol (Merck, Germany), and distilled water, respectively. ZnO NPs were first dissolved at 1% (w/v) in 70% ethanol and then used in the spin coating process. The spin coating process was performed by dropping the resulting ZnO NP solutions onto glass substrates and spun in ambient conditions for 1 min at 3000 rpm. To evaporate the solvent, the coatings were then dried on a hot plate at 90 °C for 10 min. After repeating the coating 3 times, the samples were taken to a furnace (Nabertherm, Germany) for annealing for 1 h at 400 °C, and then ZnO thin films were obtained.

Characterization of ZnO NPs and thin films

The optical property of synthesized ZnO NPs (ZnO-1 NPs and ZnO-2 NPs) and thin films (ZnO-1 thin film and ZnO-2 thin film) was characterized using a UV-Vis Spectroscopy (Thermo Evolution 3000) with a wavelength in the range of 200 nm to 800 nm, operated at a resolution of 2 nm, and maximum absorbance values were determined (Shamsuzzaman et al. 2017).

The shapes and structures of the ZnO NPs and thin films were characterized using a Hitachi S-4800 SEM-EDX device. The phase and structural information of ZnO NPs and thin films were measured by XRD analysis, which was performed with the Rigaku SmartLab 3 kW XRD system. FTIR analysis was performed to detect the presumable biomolecules responsible for the reduction of the Zn+ 2 ions and the formation and stability of ZnO NPs with an FTIR spectroscopy (Shimadzu IRAffinity-1 S HATR 10) (Al-Kordy et al. 2021).

A Krüss DSA100 model goniometer was used to investigate the hydrophobicity of ZnO thin films in the Membrane Process Engineering Laboratory, Department of Bioengineering at Manisa Celal Bayar University (Manisa, Türkiye). The equilibrium contact angle of sessile droplets was evaluated from the stabilized measurements obtained 30 s after droplet deposition on the substrate. Measurements were carried out using an Attension Optical Tensiometer, which records droplet profiles with high resolution and determines the angle through advanced curve-fitting analysis. Droplets of ultra-pure water, each with a volume of 2.5 µL, were utilized, and for each specimen, the reported contact angle corresponds to the average of three measurements taken at different surface locations following the 30-second equilibration period. To enhance statistical reliability, data were collected from at least three independent specimens, with each sample analyzed at a minimum of three distinct positions.

Antibacterial activity of ZnO NPs and thin films

Preparation of test culture and ZnO NPs suspension

In the study, Escherichia coli O157:H7 ATCC 35150 and Staphylococcus aureus 6538P were used as test cultures. The cultures were supplied from the Food Microbiology Laboratory, Food Engineering Department, Ege University, Izmir, Türkiye, and grown in Tryptic Soy Broth (TSB, pH 7.3 ± 0.2, Merck) for 24 h at 37 °C. An overnight culture of pathogens grown in TSB media at 37 °C was used to prepare the culture suspension in Phosphate Buffer Saline (PBS, 8 g/L NaCl, 1.42 g/L Na2HPO4, 245 mg/L KH2PO4, 200 mg/L KCl, pH 7.4 ± 0.2). The McFarland values of the culture suspensions were adjusted to 0.5 (≅ 8 log CFU/mL) by dilution in PBS and were used in the antibacterial tests described below.

Two different ZnO NP (ZnO-1 NPs and ZnO-2 NPs) suspensions were prepared using distilled water at a concentration of 1%. To obtain a true dispersion, the suspension was vortexed for 15 min and then subjected to ultrasonic treatment (Bandelin Sonorex) for 30 min.

Agar well diffusion method

The antibacterial activity of ZnO NPs was determined using the agar well diffusion method (Argyri et al. 2013) with some modifications. 100 µL of suspensions of test cultures was aseptically inoculated on Nutrient Agar (NA) (pH 7.0 ± 0.2, Merck) by the spread plating method, and the plates were kept for 15 min to dry in the biosafety cabinet (Thermo safe 2020). Then, 100 µL of each ZnO NP suspension was separately transferred into a hole (6 mm diameter) drilled into the agar. The plates were incubated at 37 °C for 24 h, and the antibacterial activity was measured as growth-free inhibition zones (diameter) around the well.

Minimum inhibitory concentration (MIC) and minimum bactericidial concentration (MBC)

The 96-well microplate broth dilution method was utilized to determine the MIC values of ZnO NPs according to the Clinical and Laboratory Standards Institute (Zimmer et al. 2024). Firstly, 80 µL of double-strength Mueller-Hinton Broth (MHB, pH 7.4 ± 0.2, Merck) was added to each well. Subsequently, the initial wells were loaded with 80 µL of ZnO NP suspension. Two-fold serial dilutions were then prepared, and the final concentrations of ZnO NPs were: 5 mg/mL, 2.5 mg/mL, 1.25 mg/mL, 0.625 mg/mL, 0.313 mg/mL, 0.156 mg/mL, and 0.0078 mg/mL (w/v), respectively. After dilution, 20 µL of culture suspension was added to each well. The final volume in each well was 100 µL, and then the microplates were incubated at 37 °C for 24 h. After incubation, 20 µL of 2,3,5-triphenyl tetrazolium chloride (0.5%, Merck) aqueous solution was added to the wells, and the color change of the wells was examined after 30 min incubation at 37 °C. The minimum concentration necessary for inhibition of visible growth of the test cultures (no color formation) was evaluated as the MIC value of the NPs.

To detect the MBC values, a loopful of the solution from the MIC determination was subcultured onto NA and incubated at 37 °C for 24 h. After incubation, the plates were examined for colony formation, and the lowest concentration demonstrating no visible growth was described as the MBC, indicating a 99.99% reduction in the initial inoculum because of its bactericidal effect (Kışla et al. 2024).

Direct contact test

The antibacterial efficacy of the ZnO thin films was evaluated using a direct contact test adapted from the method of Beyth et al. (2008) and Akan et al. (2023). In brief, 10 µL of bacterial suspension (106 CFU/mL) was aseptically inoculated onto the film surface and incubated at ambient temperature (25 °C) for 2 h. Bacterial recovery was performed both immediately following inoculation (t = 0) and after the 2-hour contact period (t = 2). Each specimen was subsequently homogenized in 5 mL of sterile phosphate-buffered saline (PBS) by vortexing, followed by serial 10-fold dilutions in PBS. From each dilution, 1 mL aliquots were plated on Tryptic Soy Agar (TSA) using the pour-plate technique. Plates were incubated at 37 °C for 24 h, after which colony-forming units were enumerated.

Modified Kirby-Bauer diffusion test

Test bacterial cultures were grown in Tryptic Soy Broth (TSB) at 37 °C for 24 h, after which the suspensions were standardized to a 0.5 McFarland turbidity in sterile PBS. Aliquots of 100 µL from the adjusted suspension were uniformly spread over NA plates, which were subsequently dried for 15 min under aseptic conditions in a biosafety cabinet. ZnO thin films were then carefully placed in an inverted orientation onto the agar surface and incubated at 37 °C for 24 h. Following incubation, antibacterial activity was quantified by measuring the diameter (in millimeters) of the inhibition zones formed around the test specimens, in accordance with the method described by Kışla et al. (2024).

Statistical analysis

Three independent batches were analyzed, and all experiments were conducted in three replicates. Results are presented as mean ± standard deviation. Data analysis was performed using IBM SPSS statistical software. The significance of differences (P < 0.05) was evaluated using a one-way analysis of variance (ANOVA) with Duncan’s multiple comparison module and an independent-sample t-test.

Results and discussion

Isolation and molecular identification of AAB strains used in the synthesis of ZnO NPs

In this study, fermented beverages such as vinegar [ginger vinegar (S1) and olive leaf vinegar (S2)], kombucha [produced with ginger (S3) and black mulberry (S4)], and beetroot kvass [produced with red beetroot (S5) and red beet+ginger (S6)] beverages produced by spontaneous fermentation were used as the sources of AAB. The counts of AAB found in the beverages were detected as ranging from 2.88 to 5.22 log CFU/mL (Table 1). For the isolation, cream-colored colonies with clean precipitation zones were isolated from GYC agar plates as presumptive AAB (given counts above). A total of eight presumptive AAB isolates were obtained from GYC agar. All isolates exhibited similar colony morphology, color, and growth characteristics, indicating phenotypic homogeneity. Therefore, two isolates (B1 and B2) were randomly selected as representative strains for subsequent ZnO NP synthesis and characterization. Hence, two AAB isolates, expressed as B1 and B2 strains isolated from S2 and S3 samples (respectively), were randomly selected among these isolates known as beneficial microorganisms found in the fermented beverages to synthesize ZnO NPs.

The 16 S rRNA gene sequences of B1 and B2 were aligned to NCBI sequences and shared 99.75% and 100% sequence similarity with N. hansenii strain B1 and N. hansenii strain B2, respectively. The nucleotide sequences of each strain were uploaded to GenBank in NCBI, and assigned accession numbers as PV200304 (for B1 strain) and PV200774 (for B2 strain). As expected, B1 and B2 strains had 16 S rRNA gene sequences that closely aligned to known N. hansenii ATCC type strains (Fig. 2). N. hansenii is taxonomically classified within the Acetobacteriaceae family, and it is classified into a new genus, separately from Komagateibacter genus, based on genotypic and phylogenomic analysis in 2022 (Brandão et al. 2022). Moreover, this genus was predominantly found in various fruits, vegetables, and fermented beverages produced with spontaneous fermentation. The genus is particularly known for its capacity to synthesize cellulose, a highly valued industrial material due to its natural origin, biodegradability, low toxicity, stability, and unique viscoelastic properties (Azeredo et al. 2019; Gregory et al. 2021; Thongsuk et al. 2025).

Biosynthesis of ZnO NPs by AAB

In the current study, zinc acetate aqueous solution was reduced to ZnO NPs when added to the supernatants of AAB strains identified as N. hansenii (B1 and B2). ZnO NPs formation was confirmed and characterized by various techniques. The color change into pale yellow or white precipitates showed the biotransformation of zinc ions in the presence of microbial supernatants to ZnO NPs (Rehman et al. 2019).

Several researchers reported that AAB species, such as Acetobacter xylinum can synthesize ZnO NPs by producing bacterial cellulose (Purba et al. 2024). However, to the best of our knowledge, there is no study on the synthesis of ZnO NPs using N. hansenii strains.

UV-Vis spectra of the ZnO NPs and thin films

The formation of the biosynthesized ZnO NPs and thin films was confirmed by UV-Vis spectroscopy measurements. ZnO NPs synthesized via N. hansenii B1 and B2 strains, and also thin films coated with ZnO NPs, showed their absorption peaks at 300 nm (Fig. 3a and c). The optical transmittance spectra of ZnO NPs and thin films were also presented in Fig. 3b and d, respectively. For the transmittance spectrum of nanoparticles, both films exhibited the same transmittance value (6.86%) at 433 nm. However, the optical transmittance value of ZnO-1 NPs is higher than that of ZnO-2 NPs at more than 433 nm. On the other hand, the ZnO-1 thin films showed better optical transmittance than ZnO-2 thin films in the visible range (400–800 nm). Similar values were also observed in other studies conducted, Fayed et al. (2023) reported that the UV-Vis spectroscopy absorption peak of ZnO NPs synthesized using the probiotic Bacillus coagulans appeared at 318 nm, and Elsilk et al. (2024) stated that the biosynthesized ZnO NPs by Enterobacter sp. were established via the UV-Vis absorption spectra at the wavelength range of 300 to 360 nm, which is specific for ZnO NPs. The absorption spectra of ZnO NPs observed between 265 nm and 390 nm were also reported by Mekky et al. (2021) and Vijayakumar et al. (2016).

Fig. 3.

Fig. 3

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UV-Vis absorbance spectra (a and c) and transmittance spectra (b and d) of biogenic ZnO NPs (ZnO-1 NPs synthesized via N. hansenii B1 and ZnO-2 NPs synthesized via N. hansenii B2) and ZnO thin films (ZnO-1 thin film coated with ZnO-1 NPs synthesized via N. hansenii B1; ZnO-2 thin film coated with ZnO-2 NPs synthesized via N. hansenii B2) fabricated by spin coating

SEM-EDX characterization of biosynthesized ZnO NPs and thin films

The morphology and structural characteristics of the prepared ZnO NPs were confirmed by SEM, as shown in Fig. 4. According to the SEM images, ZnO NPs were agglomerated, but the average sizes of NPs were not detected for all NPs (Fig. 4a, b). The observed agglomeration could be attributed to the presence of moisture content and interparticle interactions, including Van der Waals forces and magnetic/electrostatic attractions (Al-Dhabi et al. 2018). In the current study, the ZnO NPs exhibited a hexagonal or spherical-like morphology (Fig. 4), similar to previous findings reported by Raut et al. (2015) and Mani et al. (2022). Furthermore, their structural arrangement resembled bouquet-like formations, likely resulting from surface reconstruction within the agglomerates (Fig. 4). This phenomenon may lead to modifications in distinct crystallographic planes due to alterations in surface charges driven by active biomineralization (Pacholski et al. 2002). SEM images of ZnO-1 and ZnO-2 thin films coated on glass substrates are given in Fig. 4c and d, respectively. In SEM images, it can be clearly observed that both ZnO-1 and ZnO-2 thin films are densely, homogeneously, and uniformly coated without any needle holes or cracks. The thickness of ZnO-1 thin film was measured as 43.02 ± 6.96 μm, and the thickness of ZnO-2 thin film was measured as 427 ± 98 nm by cross-section SEM images (Fig. 5). All these results showed that the morphology of ZnO NPs was influenced by the source of the microorganism used for the synthesis.

Fig. 4.

Fig. 4

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SEM images of synthesized ZnO NPs and thin films. a ZnO-1 NPs synthesized via N. hansenii B1, b ZnO-2 NPs synthesized via N. hansenii B2, c ZnO-1 thin film coated with ZnO-1 NPs synthesized via N. hansenii B1, and d ZnO-2 thin film coated with ZnO-2 NPs synthesized via N. hansenii B2

Fig. 5.

Fig. 5

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Thickness measurements of ZnO thin films fabricated by spin coating using biogenic ZnO NPs: a ZnO-1 thin film coated with ZnO-1 NPs synthesized via N. hansenii B1 and b ZnO-2 thin film coated with ZnO-2 NPs synthesized via N. hansenii B2

EDX analysis is performed to detect the presence of metallic elements and is recognized for its ability to offer insights into the chemical analysis of the examined areas or the composition at particular spots (El-Khawaga et al. 2025). In the current study, the EDX spectra confirmed the formation of biosynthesized ZnO NPs via AAB strains, as depicted in Fig. 6, by indicating an elemental zinc (strong peak at 1 KeV) and oxygen (medium peak) peak that indicated the presence of ZnO NPs (Hamk et al. 2023). Zinc had the elemental component weight% of 44.39% for ZnO-1 NPs, 27.33% for ZnO-2 NPs while the oxygen content of 26.36%, and 27.25% respectively (Fig. 6). The other identified elemental peak signals (carbon, phosphorus, and nitrogen) from EDX analysis may be because of the broth composition (GYC) used for AAB strain growth, as well as zinc acetate used as the precursor. Similar results were observed in the previous study conducted by Al-Tameemi et al. (Al-Tameemi et al. 2023). According to the EDX spectra, the elemental zinc and oxygen peaks that indicated the presence of ZnO NPs were observed. Also, it was reported that other identified element peaks (phosphorus) signals from EDX analysis may be due to the broth composition (MRS) used for bacterial growth. Based on previous studies, ZnO NPs are impurity-free if zinc and oxygen elements are present in EDX analysis (Varadavenkatesan et al. 2019; Djearamane et al. 2022). Although EDX confirms elemental composition, accurate particle size determination typically requires techniques specifically designed for colloidal systems, such as Dynamic Light Scattering (DLS). However, for highly aggregated or biologically capped ZnO NPs, additional size-determination methods may be needed due to dispersion limitations (Singhal et al. 2011). The EDX results also confirmed the existence of zinc and oxygen elements in both ZnO-1 and ZnO-2 thin films, with the atomic percentage of 9.44% and 90.56% for ZnO-1 thin film and 7.43% and 92.57% for ZnO-2 thin film, respectively. As seen in Fig. 6, all three elements are homogeneously distributed in the structure and do not gather in a certain region in the element map. This showed that the ZnO-1 and ZnO-2 thin films are also homogeneous in elemental distribution.

Fig. 6.

Fig. 6

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EDX results of ZnO NPs and thin films: a ZnO-1 NPs synthesized via N. hansenii B1, b ZnO-1 thin film coated with ZnO-1 NPs synthesized via N. hansenii B1, c ZnO-2 NPs synthesized via N. hansenii B2, and d ZnO-2 thin film coated with ZnO-2 NPs synthesized via N. hansenii B2

XRD patterns of biosynthesized ZnO NPs and thin films

XRD analysis was performed to determine the crystallinity of the ZnO NPs, showing the states, axes, and sizes of the atoms (El-Refai et al. 2024). According to the results, ZnO NPs synthesized using supernatants of AAB strains exhibited different diffraction peaks at 2θ values in the (111) and (316) crystallographic planes, as represented in the XRD patterns of ZnO NPs (Fig. 7). These findings support the hypothesis that ZnO NPs have a monoclinic (COD database code. 4517837), and cubic (COD database code. 1537875) structures. For ZnO-1 NPs, the peaks at 2θ = 33.28° (111) and 59.39° (316) corresponded to crystallite sizes of 35.11 nm and 9.68 nm, respectively (Table 2). Similarly, for ZnO-2 NPs, peaks at 2θ = 33.28° (111) and 59.42° (316) corresponded to crystallite sizes of 26.33 nm and 9.68 nm, respectively. Moreover, the majority of the synthesized ZnO NPs exhibited a cubic ZnO crystallite type, while monoclinic Zn20O48 crystallite types were observed in ZnO-1 NPs and ZnO-2 NPs (Table 2). The size of the NPs can be used to explain the broadening of the XRD pattern, and the presence of strong peaks indicates that the product has good crystallinity (Abdul Salam et al. 2014; El-Fallal et al. 2023). On the other hand, the findings of El-Khawaga et al. (2025) and Hamk et al. (2023) reported that the XRD patterns of ZnO NPs synthesized by S. cerevisiae and B. subtilis exhibited diffraction peaks corresponding to a crystalline ZnO structure with a hexagonal wurtzite phase. XRD data of the ZnO-1 and ZnO-2 thin films coated by the spin coating technique on glass substrates are also presented in Fig. 7. No crystal peaks were observed in either ZnO thin films. These findings confirm that ZnO-1 and ZnO-2 thin films exhibited an amorphous structural configuration.

Fig. 7.

Fig. 7

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XRD results of biogenic ZnO NPs (ZnO-1 NPs synthesized via N. hansenii B1 and ZnO-2 NPs synthesized via N. hansenii B2) and thin films (ZnO-1 thin film coated with ZnO-1 NPs synthesized via N. hansenii B1; ZnO-2 thin film coated with ZnO-2 NPs synthesized via N. hansenii B2)

Table 2.

Crystallite sizes of ZnO NPs synthesized by AAB strains calculated by Scherrer equation

NPs 2θ (°) FWHM (hkl) Crystallite type Crystallite size (nm)
ZnO-1 NPs 33.28 0.23616 111 Cubic ZnO 35.11
59.39 0.94464 316 Monoclinic Zn20O48 9.68
ZnO-2 NPs 33.28 0.31488 111 Cubic ZnO 26.33
59.42 0.94464 316 Monoclinic Zn20O48 9.68
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ZnO-1 NPs synthesized via N. hansenii B1; ZnO-2 NPs synthesized via N. hansenii B2

FTIR spectra of biosynthesized ZnO NPs

FTIR was performed to validate the presence of functional groups on the surface of the synthetic materials. The spectrum of ZnO NPs is presented in Fig. 8. The observed peaks in the spectra correspond to the distinctive functional groups, which are present in the synthesized ZnO NPs (Abdelghany et al. 2023). Diverse FTIR spectra were observed between 400 and 4000 cm− 1, which facilitated the ZnO NPs by acting as reducing and capping agents (Fig. 8), which was evidence of the presence of unique functional groups of bioactive molecules bound on the surface of NPs (Farooq et al. 2022). The AAB-based ZnO NPs showed prominent peaks of 3320, 2976, 2882, 1423, 1068, 947, and 631 cm− 1 for ZnO-1 NPs, 3319, 2970, 2891, 1406, 1061, 885, 637, and 565 cm− 1 for ZnO-2 NPs (Fig. 8). The presence of –O–H groups, which were because to alcoholic, phenolic, and acid groups, was indicated by the broad peak at 3319–3350 cm− 1 (Gangwar et al. 2024). Also, the peaks at 2966, 2970, 2972, 2984, 2874, 2882, 2885, 2887, and 2891 cm− 1 are related to C–H and C -C stretching vibrations that formed because of the presence of alkane groups (Gangwar et al. 2024). Aromatic molecules were established and stated by the asymmetric stretch of C-C = C and C = C were observed at around 1406–1423 and 1057–1078 cm− 1, respectively (Raj and Lawerence 2018). The metal-oxygen (ZnO stretching vibrations) vibration pattern is correlated with the 519–631 cm− 1 absorption peak and hence 519–631 cm− 1 bands indicate the presence of ZnO NPs (Devi and Velu 2016; Doan Thi et al. 2020; Shanmugam et al. 2023).

Fig. 8.

Fig. 8

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FTIR results of biogenic ZnO NPs (ZnO-1 NPs synthesized via N. hansenii B1 and ZnO-2 NPs synthesized via N. hansenii B2)

Water contact angle (WCA) values of thin films

To assess the surface hydrophobicity of glass substrates coated with ZnO-1 and ZnO-2 thin films, a Krüss DSA100 model goniometer was employed. The WCA value of uncoated glass substrates was found to be 28.24°, while WCA values of ZnO-1 and ZnO-2 thin films coated substrates were determined as 29.56° and 25.89°, respectively (Fig. 9). The analysis indicated that coating glass substrates with ZnO-1 thin films led to a statistically significant increase in contact angle values, whereas coatings with ZnO-2 thin films resulted in a statistically significant decrease in contact angle values on the glass surfaces (P < 0.05) (Fig. 9). Matlaga et al. (1976a) noted that surfaces with contact angles in the range of 40° − 80° may exhibit strong bacterial cell adhesion, while surfaces with contact angles exceeding 65° tend to resist microbial cell attachment and can be classified as hydrophobic. The obtained WCA values clearly indicate that both ZnO-1 and ZnO-2 thin film-coated glass substrates maintain a predominantly hydrophilic surface character, as the values remain below 40°. This observation aligns with previous reports that ZnO-based coatings typically exhibit hydrophilic behavior due to the presence of surface hydroxyl groups and oxygen vacancies, which promote hydrogen bonding with water molecules (Li et al. 2022; Soltanzadeh et al. 2022). However, subtle differences in WCA between ZnO-1 and ZnO-2 thin films suggest that synthesis conditions, crystallite size, and surface roughness may influence wettability. Similar findings have been reported by other studies, where variations in NP morphology and film density significantly altered surface energy and consequently affected the hydrophilic/hydrophobic balance of ZnO-coated substrates (Boughdiri et al. 2022; Dong et al. 2022). Given the critical role of surface wettability in microbial adhesion and biofilm formation, the higher WCA value of ZnO-1 films may indicate a slightly reduced tendency for bacterial attachment compared to ZnO-2, although both coatings remain within the hydrophilic regime.

Fig. 9.

Fig. 9

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Water contact angle measurements of glass substrates before and after ZnO thin-film coating: a uncoated glass, b ZnO-1 thin film coated with ZnO-1 NPs synthesized via N. hansenii B1, and c ZnO-2 thin film coated with ZnO-2 NPs synthesized via N. hansenii B2

Antibacterial activity of biosynthesized ZnO NPs and thin films

The antibacterial activity of ZnO NPs synthesized using supernatants of AAB strains was tested both on E. coli O157:H7 ATCC 35,150 and S. aureus 6538P to represent both Gram-negative and Gram-positive bacteria, respectively. The ZnO NPs showed antibacterial effects on all the test cultures used (Table 3). The inhibition zones of ZnO NPs ranged from 12.50 to 14.00 mm, with the highest inhibitory effect on both test cultures being exhibited by ZnO-2 NPs. Similar results were reported by El-Khawaga et al. (2025), where the inhibitory effect of ZnO NPs biosynthesised by S. cerevisiae against S. aureus (22 mm) was higher than against E. coli (17 mm). In contrast to these results, ZnO NPs biosynthesized by B. subtilis ZBP4 exhibited higher inhibition zones on Gram-negative bacteria (12.00–13.30 mm), including E. coli O157:H7, than Gram-positive bacteria (11.00–12.00 mm), including S. aureus (Hamk et al. 2023).

Table 3.

Antibacterial activity of ZnO NPs synthesized by AAB strains

Inhibition diameter of ZnO NPs (mm)* MIC values of ZnO NPs (mg/mL) MBC values of ZnO NPs (mg/mL)
NPs E. coli O157:H7 S. aureus E. coli O157:H7 S. aureus E. coli O157:H7 S. aureus
ZnO-1 NPs 12.50 ± 0.71 14.00 ± 1.41 0.625 ± 0.00 1.25 ± 0.00 2.50 ± 0.00 1.25 ± 0.00
ZnO-2 NPs 14.00 ± 0.00 14.00 ± 0.00 0.625 ± 0.00 0.625 ± 0.00 1.25 ± 0.00 2.50 ± 0.00
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MIC (Minimum Inhibitory Concentration) and MBC (Minimum Bactericidal Concentration) values of ZnO-1 NPs synthesized via N. hansenii B1 and ZnO-2 NPs synthesized via N. hansenii B2 against E. coli O157:H7 and S. aureus, determined by the broth microdilution method. The concentrations of ZnO NPs ranged from 5 to 0.0078 mg/mL (w/v) in Mueller-Hinton Broth. Microplates were incubated at 37 °C for 24 h, and MIC values were determined using TTC as a viability indicator. MBC values were assessed by subculturing onto nutrient agar followed by incubation at 37 °C for 24 h. *Well diameter was not included

The MIC values of the ZnO NPs varied in the range of 0.625 to 1.25 mg/mL for both pathogenic bacteria. Also, MIC values showed that ZnO-1 NPs were more effective against E. coli O157:H7 than S. aureus, in parallel with inhibition zone values (Table 3). In addition, ZnO NPs exhibited high bactericidal activity at concentrations ranging from 1.25 to 2.50 mg/mL. The antibacterial activity of ZnO NPs can show differences depending on the sources of NPs and microorganisms tested. In this scope, in a study, it was reported that the MIC values of ZnO NPs biosynthesized by B. haynesii for E. coli and S. aureus were determined as 8 and 4 mg/mL, respectively (Rehman et al. 2019); also, it is seen that these values, compared to our results, were higher than the MIC values of both ZnO NPs synthesized by AAB on E. coli and S. aureus. Additionally, in the study conducted by Elsilk et al. (2024), in which ZnO NPs mediated by Enterobacter sp. showed antimicrobial activity against E. coli, S. typhimurium, Klebsiella pneumoniae, and Candida albicans with different MIC values at 0.4, 0.9, 1.0, and 1.5 mg/mL, respectively.

The results of the modified Kirby-Bauer test, conducted to evaluate the antimicrobial activity of glass substrates coated with ZnO NPs against pathogenic E. coli O157:H7 and S. aureus, are presented in Fig. 10. Analysis indicated that the highest inhibition zone (2.38 mm) was observed on ZnO-2 thin films against E. coli O157:H7 while the lowest was on ZnO-2 thin film against S. aureus. The inhibition zones of ZnO-1 thin film were found to be 1.00 mm and 1.75 mm against both E. coli O157:H7 and S. aureus. The inhibition zones of ZnO-2 thin film were measured as 2.38 mm for E. coli, and 0.50 mm for S. aureus, respectively. In the direct contact tests, both E. coli O157:H7 and S. aureus, at a concentration of approximately 104 cfu/cm2, were inoculated onto 1 cm × 1 cm glass substrates (ZnO NPs-coated and uncoated). Following inoculation, samples were cultured immediately (t: 0 h) and after a 2-hour direct contact period (t: 2 h) at room temperature. Additionally, the inhibition rates of microbial cells following a two-hour contact period with ZnO NPs-coated surfaces were calculated using the formula:

Fig. 10.

Fig. 10

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Modified Kirby-Bauer results of thin films coated on glass substrates. Antibacterial performance of ZnO thin-film coatings (ZnO-1 thin film coated with ZnO-1 NPs synthesized via N. hansenii B1; ZnO-2 thin film coated with ZnO-2 NPs synthesized via N. hansenii B2) assessed against E. coli O157:H7 and S. aureus. Bacterial suspensions were standardized to 0.5 McFarland, spread on nutrient agar plates, and ZnO thin films were placed onto the agar surface. Plates were incubated at 37 °C for 24 h, and antibacterial activity was quantified by measuring inhibition zone diameters (mm). In the figure, different lower-case letters (a, b) in the same row and different capital letters (A, B) in the same column on the data table indicate a significant difference (P < 0.05) between mean values

graphic file with name d33e1585.gif

Where Nt:0 represents the initial inoculated cell count (CFU/cm2), and Nt:2 denotes the microbial cell count (CFU/cm2) measured after two hours of contact with the ZnO NPs-coated surfaces. The data obtained from the analysis are presented in Table 4. As seen in Table 4, E. coli O157:H7 was completely inhibited by both ZnO-1 and ZnO-2 thin films, while S. aureus was totally inhibited by only ZnO-1 samples. On the other hand, the number of S. aureus significantly reduced (1.16 log units) after two hours of direct contact with ZnO-2 thin films.

Table 4.

The viable cell counts of various microorganisms following direct contact time at 0 (t: 0) and 2nd (t: 2) hours on the glass substrates coated with the ZnO NPs

Thin Films E. coli O157:H7 (log CFU/cm2) S. aureus (log CFU/cm2)
t: 0 t: 2 t: 0 t: 2
ZnO-1 Thin Film 4.02 ± 0.15A, b* < 0.7a 4.15 ± 0.11A, b < 0.7a
ZnO-2 Thin Film 3.96 ± 0.31A, b < 0.7a 4.57 ± 0.41A, c 3.41 ± 0.57b
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Antibacterial activity of ZnO thin-film coatings (ZnO-1 thin film coated with ZnO-1 NPs synthesized via N. hansenii B1; ZnO-2 thin film coated with ZnO-2 NPs synthesized via N. hansenii B2) was evaluated by the direct contact test against E. coli O157:H7 and S. aureus. A bacterial suspension (10 µL, 106 CFU/mL) was inoculated onto the film surface and incubated at 25 °C for 2 h. Bacterial recovery was performed at t: 0 and t: 2 h, followed by plate counting after incubation at 37 °C for 24 h. *Mean values in the same row that are not followed by the same lower-case letter (a-b) are significantly different (P < 0.05). Mean values in the same column that are not followed by the same capital letter (A-D) are significantly different (P < 0.05). The limit of detection is 0.7 log CFU/cm2 for direct contact analysis of ZnO thin films

The differences observed in the inhibition zones and direct contact assays between ZnO-1 and ZnO-2 thin films may be attributed to the variations in the particle size, crystallinity, and surface area, which strongly influence antimicrobial performance (Stanković et al. 2013). Previous studies have emphasized that smaller ZnO nanoparticles with higher surface-to-volume ratios can interact more efficiently with bacterial membranes, enhancing their bactericidal activity (De Souza et al. 2019). The complete inhibition of E. coli O157:H7 by both thin films suggests that Gram-negative bacteria may be more susceptible to ZnO-mediated antibacterial mechanisms, likely due to the thinner peptidoglycan layer in their cell walls. In contrast, the reduced inhibition observed against S. aureus is consistent with the greater structural resistance of Gram-positive bacteria, which possess a thicker peptidoglycan barrier (Malik et al. 2022). The antibacterial action of ZnO coatings has been associated not only with physical interactions and membrane disruption but also with the generation of reactive oxygen species (ROS), such as H2O2, which can penetrate bacterial cells and damage essential biomolecules (Xie et al. 2011). These findings highlight the multifactorial antimicrobial mechanisms of ZnO thin films and confirm their potential for applications in antimicrobial coatings.

The antibacterial mechanism of ZnO NPs likely involves their accumulation within the cytoplasm or outer membrane of bacterial cells, facilitating the release of Zn2+ ions. This process leads to the disruption of bacterial cell membranes, structural damage to membrane-associated proteins, and genomic instability, ultimately resulting in bacterial cell death (Qais et al. 2019; Wang et al. 2019). Studies have demonstrated that ZnO NPs exert their bactericidal activity through direct interaction with the phospholipid bilayer, compromising membrane integrity. Notably, the antimicrobial efficacy of ZnO NPs was significantly diminished upon the introduction of radical scavengers such as glutathione, vitamin E, and mannitol, suggesting that reactive oxygen species (ROS) generation plays a pivotal role in their antibacterial properties. However, the antimicrobial activity did not appear to be solely attributable to Zn2+ ions released from ZnO NP suspensions. Evaluation of ZnO NPs against various microorganisms revealed variations in their MIC values depending on antibiotic resistance profiles, indicating potent antibacterial activity. These findings suggest that the release of Zn2+ ions from ZnO NPs substantially contributes to their bactericidal effect (Elsilk et al. 2024; Gökmen et al. 2024). On the other hand, when the antimicrobial efficacy was evaluated in relation to the WCA results, although the WCA values of ZnO-1 and ZnO-2 thin films differ only slightly (approximately 3–4⁰), both surfaces remain within the hydrophilic regime (< 40⁰), where strong bacterial adhesion is generally expected rather than inhibited (Matlaga et al. 1976). Previous studies have indicated that pronounced reductions in bacterial attachment are typically associated with more substantial shifts toward hydrophobicity (WCA > 65⁰) (Matlaga et al. 1976b; Wang et al. 2012; Gao and Guo 2017). Therefore, the observed differences in antibacterial performance between ZnO-1 and ZnO-2 thin films cannot be attributed solely to surface wettability. Nevertheless, even minor variations in WCA may influence the initial stages of bacterial attachment under short contact conditions, acting as a secondary factor alongside dominant mechanisms such as nanoparticle size, surface area, crystallinity, ROS generation, and Zn2+ ion release (Akan et al. 2023). In this context, the slightly higher WCA of ZnO-1 thin films may contribute marginally to reduced bacterial adhesion; however, the antibacterial activity of the ZnO thin films should be considered the result of multifactorial surface and bacteria interactions rather than wettability alone.

Although ZnO is generally GRAS and widely used in food-related applications, safety considerations for food-contact materials require careful evaluation beyond antibacterial efficacy. A comprehensive safety assessment of nanoparticle-based food-contact materials remains essential, as nanoparticles may enter the human body through ingestion, inhalation, or dermal exposure, with potential accumulation due to their limited solubility in biological fluids (Maisanaba et al. 2015). For food packaging applications, the migration of ZnO nanoparticles or released Zn2+ ions into food matrices is a critical concern and is influenced by factors such as particle size, concentration, temperature, food composition, and pH (Huang et al. 2015). Previous studies have reported that ZnO nanoparticles may induce cytotoxic or genotoxic effects in vitro, including oxidative stress and reduced viability in human epidermal and intestinal (Caco-2) cells at certain concentrations (Sharma et al. 2009; Mao et al. 2016), although protective effects of antioxidants have also been observed (Wu et al. 2019). Current toxicological evidence is largely derived from in vitro or animal models, highlighting the need for cautious interpretation (McCracken et al. 2016). Regulatory frameworks established by the European Food Safety Authority (EFSA) and the FDA emphasize that food-contact materials must not release constituents at levels posing risks to human health (EU 2004; FDA 2018). Therefore, future work should focus on nanoparticle migration, coating stability, and cytotoxicity evaluations to support the safe application of ZnO-based thin films in food packaging.

Conclusion

In this study, ZnO NPs were synthesized through an eco-friendly approach using extracellular extracts of AAB strains derived from fermented foods, and thin films with ZnO NPs were successfully fabricated via spin coating techniques. N. hansenii strain B1 and N. hansenii strain B2 were isolated from fermented beverages and used for the synthesis of ZnO NPs. The UV-Vis spectrum confirmed the extracellular synthesis of ZnO NPs and the formation of thin films coated with ZnO NPs. SEM analysis showed that ZnO NPs were agglomerated and homogeneous nanosized thin films, while EDX confirmed their pure phases. XRD analysis revealed monoclinic and cubic crystalline structures with crystallite sizes ranging from 9.68 to 35.11 nm, while thin films showed noncrystalline structures. FTIR spectra exhibited characteristic peaks, further validating the structural properties of the NPs. Additionally, thin films were evaluated as hydrophilic, showing distinct contact angles of 29.56° for ZnO-1 and 25.89° for ZnO-2. Furthermore, ZnO NPs and thin films exhibited strong antibacterial activity against E. coli O157:H7 and S. aureus. Based on these results, ZnO NPs synthesized by AAB isolated from fermented beverages via a green synthesis approach can be incorporated as antimicrobial agents into packaging materials for the food industry and related fields. Additionally, thin films produced from these NPs can be used as antimicrobial surfaces in food industry applications. Nevertheless, certain limitations of the present study should be acknowledged. Although ZnO-based materials are widely considered safe, the absence of zinc ion migration/leaching analyses, long-term coating durability studies, and cytotoxicity assessments limits direct claims regarding food-contact and biomedical applications. Future research will therefore focus on evaluating Zn2+ release under relevant conditions, assessing coating stability and wear resistance, and conducting comprehensive biocompatibility and antibiofilm studies to ensure safe and reliable real-world use.

Supplementary Information

Below is the link to the electronic supplementary material.

Supplementary Material 1 (72.5KB, docx)

Acknowledgements

The author gratefully thanks to Dr. Gökhan Gurur Gökmen for his support.

Funding

Open access funding provided by the Scientific and Technological Research Council of Türkiye (TÜBİTAK).

Data availability

The datasets generated during and/or analyzed during the current study are available from the corresponding author on reasonable request.

Declarations

Conflict of interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this article.

Ethical approval

Working only in bactericidal culture and this work does not contain any human/animal sample.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary Material 1 (72.5KB, docx)

Data Availability Statement

The datasets generated during and/or analyzed during the current study are available from the corresponding author on reasonable request.

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