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. 2025 Jul 20;34(15):3739–3751. doi: 10.1007/s10068-025-01949-1

Evaluation of the antimicrobial and antibiofilm properties of Quercus phillyraeoides essential oil against Staphylococcus aureus

Ye-Seul Yun 1,#, Soo-Hwan Kim 2,#, Sang-Hyun Park 1,✉
PMCID: PMC12528580  PMID: 41113255

Abstract

This study investigated the antimicrobial activity and underlying inactivation mechanisms of Quercus phillyraeoides essential oil (QP-EO) against Staphylococcus aureus. Results of the disk diffusion assay demonstrated the potent antimicrobial activity of QP-EO (zone of inhibition, 11.97–12.08 mm). The minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC) of QP-EO against S. aureus was 1.25 mg/mL, indicating its potential bactericidal properties. This similar MIC and MBC values suggests that QP-EO induces irreversible damage at specific concentrations. In addition, transmission electron microscopy revealed that QP-EO induces cell shape distortion and cell wall rupture in S. aureus, which may be caused by the small essential oil particles. QP-EO also significantly inhibited S. aureus initial biofilm formation and reduced the cell mass of preformed biofilms, suggesting its potential in controlling biofilm-associated infections. These findings highlight the efficacy of QP-EO as a natural antimicrobial and antibiofilm agent, with implications for microbial control and food safety.

Supplementary Information

The online version contains supplementary material available at 10.1007/s10068-025-01949-1.

Keywords: Staphylococcus aureus, Quercus phillyraeoides essential oil, Antimicrobial property, Inactivation mechanism, Biofilm inactivation

Introduction

Foodborne pathogens cause a multitude of intestinal diseases, resulting in significant economic losses, and are a major public health concern. Despite advancements in inactivation technologies and the implementation of Hazard Analysis and Critical Control Point (HACCP), approximately 9 million cases of illnesses, 56,000 hospitalizations, and 1,300 deaths attributed to foodborne pathogens have been reported annually (White et al., 2022). Staphylococcus aureus has emerged as a representative gram-positive pathogen not only in the food industry but also in the medical field for decades. S. aureus infections range from food poisoning and mild skin infections to potentially fatal conditions such as bacteremia, endocarditis, necrotizing pneumonia, and toxic shock syndrome. Staphylococcal enterotoxin type A (SEA) is responsible for food poisoning and emetic syndrome, whereas other staphylococcal toxins such as superantigen and toxic shock syndrome toxin-1 are not involved in emesis (Suzuki et al., 2020). Considering that the sea gene is generally expressed in the mid-exponential phase, the growth of S. aureus should be inhibited to prevent staphylococcal poisoning (Homsombat et al., 2021). However, once S. aureus adhere to surfaces, biofilm could be formed and make challenge to inactivate the bacteria. Biofilm formation initiates with the attachment of single cells to the surface and the formation of cell clusters. Then, for maturation, secretion of extracellular polymeric substances (EPS) occurs to encase the cells. Previous articles have demonstrated that the complex three-dimensional structure and EPS of biofilms confer the enhanced physical and chemical resistance (Kim et al., 2022).

Essential oils are concentrated extracts derived from various plant parts and have been extensively utilized in diverse fields, including therapeutics, pharmaceutical ingredients, and food additives, owing to their therapeutic properties and bioactive components (Ali et al., 2015). The constituents of essential oils are commonly divided into terpenes, which are composed of various amounts of isoprene, and other hydrocarbons such as aliphatic and aromatic hydrocarbons. Some terpenes and terpenoids (Souza et al., 2022), as well as aromatic hydrocarbons (da Silva et al., 2023) exhibited antimicrobial activity. Owing to their antimicrobial properties, essential oils have been utilized as natural preservatives or in active packaging (Carson et al., 2002). Although their unique odor and flavor generally pose limitations to their application in the food industry, the appropriate utilization of essential oils can improve the organoleptic properties of food. Therefore, there is a continuous demand for the development and study of various essential oils.

The genus Quercus, commonly referred to as oak, has a long history of use in traditional medicine worldwide owing to its antiseptic properties and therapeutic effects on gastroenteritis (Taib et al., 2020). Quercus phillyraeoides, which is widely distributed in subtropical areas, has been studied for its potential as an antioxidant and antidiabetic medication (Indrianingsih and Tachibana, 2016). However, to the best of our knowledge, the antimicrobial activity of Q. phillyraeoides has not been evaluated. In this study, we aimed to evaluate the antimicrobial property of Q. phillyraeoides essential oil (QP-EO) against S. aureus and investigate the underlying inactivation mechanisms.

Materials and methods

Bacterial cultures and cell suspension

Three strains of S. aureus (ATCC 25923, ATCC 27213, and ATCC 29213) were obtained from the bacterial culture collection of the Department of Food Science and Biotechnology, Seoul National University (Seoul, South Korea). Stock solution of each S. aureus strain was maintained at −80°C in 70% glycerol. To prepare the working solution, each strain was streaked onto tryptic soy agar (TSA; MB cell, Seoul, Korea), incubated at 37°C for 24 h, and then stored at 4°C. Subsequently, a single colony was inoculated into 5 mL of tryptic soy broth (TSB; MB cell) and incubated at 37°C for 24 h. Bacterial cells were collected via centrifugation at 4000 × g for 15 min at 4°C, and washed three times with sterile phosphate-buffered saline (PBS; LPS solution, Daejeon, Korea). The pellets were then resuspended in 0.2% peptone water (PW; MB cells), corresponding to approximately 106–107 CFU/mL.

Evaluation of antimicrobial activity

QP-EO dissolved in ethanol obtained from The Korea Plant Extract Bank at the Korea Research Institute of Bioscience and Biotechnology (Daejeon, Korea). QP-EO concentrations at 10, 5, 2.5, 1.25, 0.625, and 0.3125 mg/mL were prepared using TSB. The zone of inhibition (ZOI), minimal inhibitory concentration (MIC), and minimal bactericidal concentration (MBC) were determined to assess the antimicrobial activity of QP-EO.

Disc diffusion assay

Each S. aureus strain (0.1 mL) was spread on TSA and allowed to dry for 10 min. Then, an 8-mm sterile filter paper (Advantec, Tokyo, Japan) was placed on the inoculated TSA and inoculated with 10 μL of 10 mg/mL QP-EO. The plates were incubated for 24 h at 37°C, and the ZOI (mm) was measured.

MIC and MBC assay

Concentrations of QP-EO ranging from 0.3125 to 10 mg/mL was prepared via serial two-fold dilution in TSB. Each well of a 96-well microtiter plate (SPL Life Sciences, Pocheon, Korea) was inoculated with 100 μL of each QP-EO concentration and 100 μL of S. aureus cell suspension (5 log CFU/mL), and subsequently incubated for 24 h at 37°C. After incubation, the absorbance of each well was measured at 600 nm using a spectrofluorophotometer (SpectraMax iD3; Molecular Devices, CA, USA). The MIC was determined at an optical density of < 0.05.

To determine the MBC of QP-EO, 100 μL of each S. aureus strain was treated with QP-EO concentrations below the MIC. Each treated strain was then spread-plated on TSA and incubated for 24 h at 37°C. The MBC was determined when more than three log reductions were achieved.

Time-kill analysis

Culture cocktails containing the three S. aureus strains were inoculated (final concentration, 106 CFU/mL) in TSB supplemented with the MIC of QP-EO. To determine bacterial viability, aliquots were taken at 1, 3, 6, 12, and 24 h, diluted in 0.2% PW, and plated onto TSA. The plates were incubated for 24 h at 37°C, and colonies were counted after incubation.

Growth curve analysis

Each strain of S. aureus was diluted to 106 CFU/mL in TSB. One hundred microliters each of the culture suspension and QP-EO (MIC, 1/2 MIC, 1/4 MIC, 1/8 MIC) were added into each well in a 96-well microtiter plate. The control well was inoculated with 200 μL of culture suspension. The 96-well plate was incubated for 24 h at 37°C, and cell growth was measured every hour at 600 nm using a spectrofluorophotometer.

Analysis of the mechanisms underlying antimicrobial activity

Fluorescence analysis was conducted to investigate the underlying inactivation mechanisms. Briefly, S. aureus cells were treated with 1.25 mg/mL QP-EO (MIC and MBC) for 1 h. Treated or untreated samples were then transferred to a FILTRA-BAG (Labplas, Quebec, Canada) and filtered using a stomacher to eliminate the polymeric substances in QP-EO. Filtered aliquots were centrifuged at 4,000 × g for 20 min at 4°C. The supernatant was used for measuring the extracellular DNA and protein, whereas bacterial pellets were used for assessing membrane damage, intracellular ROS content, and respiratory enzyme activity. Pellets were washed thrice using PBS.

Membrane damage assessment

Propidium iodide (PI; Sigma-Aldrich, St. Louis, USA) uptake test and DiBAC4(3) staining were performed to assess membrane integrity and potential in S. aureus after QP-EO treatment, respectively. Briefly, bacterial cell pellets were resuspended in PBS and added PI or DiBAC4(3) solution to be final concentration of 2.9 μM or 4.8 mM, followed by incubation for 10 or 15 min at 37°C in the dark, respectively. The cells were collected via centrifugation at 10,000 × g for 1 min, washed twice with PBS, and resuspended in 1 mL PBS. For measuring fluorescence values of PI uptake test or DiBAC4(3) staining, excitation wavelength of 493 nm or 488 nm and emission wavelength of 630 nm or 525 nm was utilized with a spectrofluorophotometer. The fluorescence values were normalized by OD600.

Extracellular DNA and protein measurement

The supernatant extracted from treated or untreated sample was filtered through a 0.45 µm syringe filter (Merck Millipore, MA, USA) to removed residual cells and macromolecules. Subsequently, the ultraviolet absorbance was measured at 260 and 280 nm using a spectrofluorophotometer.

Intracellular ROS (iROS) concentration and respiratory enzyme activity measurement

The iROS concentration and respiratory enzyme activity were measured using CM-H2DCFDA (Invitrogen, MA, USA) and iodonitrotetrazolium chloride (INT; Sigma-Aldrich), respectively. Bacterial cell pellets were resuspended in 5 μM CM-H2DCFDA or 80 μM INT solution and then incubated for 15 min or 2 h, respectively, at 37°C in the dark. The suspension was centrifuged at 10,000 × g for 1 min, and the cell pellets were collected and washed twice with PBS. CM-H2DCFDA-treated sample were resuspended in 1 mL PBS. Meanwhile, the INT-treated sample was dissolved in 1 mL of 1:1 (v/v) ethanol: acetone and then incubated for 10 min at 37°C to extract iodonitrotetrazolium formazan (INF). Fluorescence values were measured at excitation and emission wavelengths of 495 and 525 nm, respectively, and the absorbance at 490 nm was measured to quantify INF. The fluorescence and absorbance values were normalized to OD600.

Transmission electron microscopy (TEM)

Specimen preparation included modified microwave-assisted post-fixation, dehydration, resin infiltration, and polymerization methods (Pelco BioWave Pro; Ted Pella, Inc., CA, USA). Treated or untreated S. aureus strains were centrifuged at 4,000 × g for 20 min at 4°C. Cell pellets were then collected, washed three times with PBS, resuspended in modified Karnovsky’s fixative containing 2% paraformaldehyde and 2% glutaraldehyde in 0.05 M sodium cacodylate buffer (pH 7.2), and finally incubated overnight at 4°C. After primary fixation, each sample was centrifuged at 10,000 × g for 1 min and washed three times with 0.05 M sodium cacodylate buffer (pH 7.2) for 5 min. The cells were resuspended in 1% osmium tetroxide in 0.05 M sodium cacodylate buffer (pH 7.2) for post-fixation, microwaved at 35°C for 40 s, and washed three times with distilled water at 25°C. Then, the cells were stained with 0.5% uranyl acetate, microwaved at 35°C for 40 s, and incubated at 4°C for 2 h. The cells were dehydrated serially using a graded ethanol (30, 50, 60, 70, 95, and 100%), microwaved for 40 s, and washed three times with 100% ethanol. Transition was conducted with 100% propylene oxide, followed by microwaving at 30°C for 40 s. For infiltration, the cells were immersed in a 1:1 and 1:2 solution of propylene oxide and Spurr’s resin solutions, microwaved at 30°C for 15 min, and then replaced twice with Spurr’s resin with microwaving for 15 min. To obtain specimen blocks, the resin was transferred to an oven at 70°C for 24 h for polymerization. The specimens were then sliced (70-nm thick) using an ultramicrotome (MT-X; RMC, Tucson, AZ, USA) and stained with 2% uranyl acetate for 10 min, followed by Reynolds’ lead citrate for 10 min. The sections were observed using an 80-kV transmission electron microscope (JEM1010; ZEOL, Japan).

Inhibition of biofilm formation and matured biofilm

To assess inhibition ability of QP-EO, the QP-EO was serially diluted in TSB to 1/2 MIC (0.625 mg/mL) and 1/4 MIC (0.3125 mg/mL), and the bacterial suspension was diluted to a concentration of 6 log CFU/mL. The Diluted QP-EO 100 µL and bacterial suspension 100 µL were mixed in a 96-well plate. The positive control was prepared by adding 100 µL of TSB to the bacterial suspension instead of the extract, and the negative control was prepared by adding TSB without the bacterial suspension. The 96-well plate was sealed and incubated at 37°C for 48 h.

For biofilm formation, 200 µL of bacterial suspension corresponding to 6 log CFU/mL, was added to a 96-well plate and incubated at 37°C for 24 h. To maintain cell viability, after 24 h, the TSB in each well was replaced with fresh TSB, and the plate was incubated again at 37°C for 24 h. After a total of 48 h of incubation, the supernatant was removed and the wells were washed once with PBS. Subsequently, 200 µL of QP-EO, serially diluted in TSB to be 2 MIC (2.5 mg/mL), MIC (1.25 mg/mL), and 1/2 MIC (0.625 mg/mL), was added to the wells, and the plate was incubated at 37°C for 0, 6, and 12 h.

Biofilm quantification

To measure the total biomass content, and the polysaccharide and protein secretion of S. aureus biofilm, crystal violet assay, calcofluor white staining, and fluorescein isothiocyanate (FITC) staining were performed. After QP-EO treatment, the extract was removed from the each well and washed once with sterile DW to remove non-adherent cells or EPS. Then, the subsequent staining was performed as follows for each:

Crystal violet assay (Total biomass content)

After washing, the biofilm was fixed at 60°C for 60 min. Each well was stained with 125 µL of 0.1% crystal violet (Samchun Chemicals, Seoul, Korea) solution and incubated for 20 min at 37°C. The unbound crystal violet was removed and each well washed once with sterile DW. Then, 200 µL of 30% acetic acid (Daejung Chemicals & Metals Co., Ltd, Siheung, Korea) was added to each well and incubated at 4°C for 20 min to dissolve the crystal violet bound to the biofilm. The absorbance at 570 nm was measured using a SpectraMax iD3 (Molecular Devices) to quantify biofilm formation. To determine viable cell counts, the biofilm was scraped off the well surface using a sterile loop (SPL Life Sciences), and 100 µL of the cell suspension was serially diluted tenfold in PW to 900 µL and spread on TSA. The plates were incubated at 37°C for 24 h, and colonies were counted and expressed as log CFU/mL.

Total biomass content%=Absorbance of control-Absorbance of treated cells/Absorbance of control×100

Calcofluor white staining (Polysaccharide content)

After washing, each well was stained with 250 µL of 1 mg/mL calcofluor white staining solution (Sigma-Aldrich) and incubated in the dark at room temperature for 30 min. The solution was then removed and the wells were washed once with PBS. Calcofluor white was dissolved with 200 µL of 99.9% EtOH (Daejung Chemicals & Metals Co., Ltd) and measured using a SpectraMax iD3 (Molecular Devices) with an excitation wavelength of 360 nm and an emission wavelength of 460 nm.

Polysaccharide content%=Fluorescence value of control-Fluorescence value of treated cells/Fluorescence value of control×100

FITC staining (protein content)

After washing, each well was stained with 250 µL of 20 µg/mL fluorescein 5(6)-isothiocyanate (FITC, MilliporeSigma, USA) solution dissolved in dH2O was added and incubated in the dark at room temperature for 30 min. The solution was then removed and the wells were washed once with PBS. FITC in biofilm was dissolved with 200 µL of dH2O and measured using a SpectraMax iD3 (Molecular Devices) with an excitation wavelength of 485 nm and an emission wavelength of 528 nm.

Protein content%=Fluorescence value of control-Fluorescence value of treated cells/Fluorescence value of control×100

Statistical analysis

All experiments were performed in triplicate. Data are expressed as means ± standard deviations (SD). Statistical analysis was performed using analysis of variance (ANOVA) and Duncan’s multiple range test in the Statistical Analysis System (SAS Institute, Cary, NC, USA). Differences with p < 0.05 were considered statistically significant.

Results and discussion

Although the antimicrobial activities of essential oils have been substantially elucidated, many phytochemicals remain underexplored. To the best of our knowledge, the present study is the first to explore the antimicrobial properties of QP-EO and its inactivation kinetics against S. aureus. Furthermore, the inactivation mechanism of QP-EO was elucidated using fluorescence spectrophotometry and TEM image analyses.

Antimicrobial properties of QP-EO

A previous study revealed that leaves of Quercus spp. contain antimicrobial agents such as β-sitosterol (Indrianingsih and Tachibana, 2016). In addition, various phenolic compounds and flavonoids that exhibit potential antimicrobial properties have been reported as components of Quercus species (Kim et al., 2012). One of the most commonly used methods to observe the inhibition of microbial growth by antimicrobial agents is the disk diffusion assay. This method involves placing the antimicrobial agent on a disk; when placed near the bacteria, bacterial growth is inhibited, leading to the formation of a clear zone around the disk. The diameter of this clear zone was measured to evaluate antimicrobial potency. In this study, the ZOI produced by QP-EO against S. aureus strains ranged from 11.97 to 12.08 mm (Table S1). A clear zone was evidently observed around the QP-EO-coated disk, as shown in Fig. S1. Subsequently, the MIC and MBC of QP-EO against S. aureus was determined as 1.25 mg/mL. Shariatifar et al. (2014) reported similar MIC for Quercus infectoria leaves extracts of 1.024 mg/mL against S. aureus. The MBC of lavender oil against S. aureus has been reported to range from 1.56 to 12.5 mg/mL. In comparison, the MBC of QP-EO observed in this study was lower, indicating its ability to control S. aureus at a comparatively lower concentration than some essential oils (Truong and Mudgil, 2023). Interestingly, the MIC and MBC values of QP-EO against S. aureus were similar, indicating that the concentration of QP-EO required to inhibit the growth of S. aureus was sufficient to kill the bacteria. Although fine adjustments to QP-EO concentration may allow for a difference between the MIC and MBC, it is improbable for the difference to exceed 0.6 mg/mL. Kot et al. (2019) reported that the MBC/MIC ratio varies widely, ranging from 1 to > 10, depending on the composition of the essential oil. Based on this evidence, we hypothesized that QP-EO exerts its antimicrobial effects by causing irreversible damage to microorganisms at certain concentrations.

The growth curves of the three S. aureus strains treated with QP-EO (MIC, 1/8, 1/4, 1/2 MIC) are shown in Fig. 1A. In the control group, the exponential phase in the three strains initiated after 4 h. However, when treated with 1/8 and 1/4 of MIC, the exponential phase initiated after 6 h, and with 1/2 of MIC after 8–10 h. Stationary phase was initiated after 14–16 h in the control group, with OD readings of 0.2–0.3, and at least 20 h in the 1/2 MIC group (OD 0.15–0.2). Staphylococcal toxins are produced during the exponential phase and are influenced by environmental factors such as temperature (Abolghait et al., 2020). Several studies have shown that the agr quorum sensing system, which is regulated by cell density, is involved in staphylococcal toxin production (Todd et al., 2019). Thus, delaying the time required to reach the exponential growth phase or preventing an increase in cell density may reduce the possibility of staphylococcal food poisoning.

Fig. 1.

Fig. 1

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The effect of Q. phillyraeoides EO (control, 1/8 MIC, 1/4 MIC, 1/2 MIC and MIC) on the growth of S. aureus ATCC 25923, 27213, 29213 (A), and time-kill kinetics of S. aureus treated with 1.25 mg/mL Q. phillyraeoides EO (B). Each point and set of error bars represent the mean value for three replicates and the standard deviation, respectively

Based on the time-kill kinetics (Fig. 1B), QP-EO treatment using the MBC exhibited a prolonged tailing effect. A 2.44-, 3.2-, and 3.79-log reduction were observed after 1, 6, and 24 h of QP-EO treatment, respectively. Basri and Khairon (2012) reported that a tailing effect occurs after 6–8 h of Quercus infectoria galls extract treatment at 1/8 MIC. In addition, diverse antimicrobial constituents within essential oils can induce variations in inactivation kinetics, and tailing effects have been observed at the MIC in most cases (Carson et al., 2002). To overcome this limitation, previous studies have attempted to combine essential oils with other essential oils or detergents (Kwiatkowski et al., 2019), Thus, further studies should be conducted to enhance the microbicidal effects of QP-EO to achieve more than a 5-log reduction.

Inactivation mechanisms of Q. phillyraeoides EO

Previous studies have attempted to shed light on the inactivation mechanisms of various essential oils (Niu et al., 2019). Phenols, aldehydes, terpenes, alcohols, ethers, and ketones play pivotal roles in the antimicrobial effects of these essential oils (Dhakad et al., 2018). In this study, PI uptake and DiBAC4(3) accumulation increased in S. aureus after QP-EO treatment at MIC for 1 h (Fig. 2A). An increase in PI uptake indicates that the antimicrobial agent induces cell membrane dysfunction. PI can intercalate into the major groove of dsDNA, but cannot penetrate intact cell membranes (Hare and Johnston, 2011). Meanwhile, DiBAC4(3) enters the cell membrane when the membrane potential changes, resulting in an enhanced fluorescence (Zhao et al., 2019). Traditionally, DiBAC4(3) dye has been used to distinguish between live and dead cells based on the loss of membrane potential in dead cells. Nevertheless, it has been reported that electroheating phenomena such as ohmic heating can also lead to a reduction in the membrane potential (Kim and Kang, 2017). Previous studies have observed this type of cell membrane damage by essential oils. For instance, 1.0 mM thymol and cymene reduced the number of B. cereus vegetative cells by 2–3 log CFU/mL, showing a tailing effect after 20 min (Delgado et al., 2004).

Fig. 2.

Fig. 2

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Propidium iodide (PI) uptake value and DiBAC4(3) accumulation value (A), Extracellular DNA content and protein content (B), and ROS and respiratory chain dehydrogenease activity value (C) of S. aureus after treatment with 1.25 mg/mL Q. phillyraeoides EO for 1 h. Different uppercase letters indicate significant (p < 0.05) differences

DNA and proteins are the cardinal components of cells that determine viability, and their leakage indicates that the cell membrane has undergone severe damage (Song et al., 2016). In this study, MIC and MBC values of QP-EO against S. aureus, coupled with the irreversible disruption of the cell membrane (Fig. 2B), suggesting that QP-EO may lead to bacterial death before growth can be achieved. Hu et al. (2019) reported that Listea cubeba essential oil (LC-EO), which contains citral, disrupts the cell membrane of S. aureus, leading to leakage of DNA and proteins. Notably, they also showed that bacterial death was observed at the MIC of LC-EO, similar to the findings in the present study. Liu et al. (2023) demonstrated that 1 mg/mL eugenol, which is present in Querecus spp., increases the cell permeability of Klebsiella pneumoniae, resulting in the leakage of bacterial cytoplasmic components. Moreover, they reported that the intracellular ROS (iROS) levels increased after eugenol treatment, leading to ROS-induced membrane rupture. In the present study, we also measured the iROS levels in S. aureus to determine whether cell membrane disruption could be attributed to ROS accumulation. However, we did not observe protein denaturation or an increase in oxidative stress within the cells (Fig. 2C).

Irreversible membrane disruption is not always observed when essential oils cause leakage of cell components. Hyldgaard et al. (2015) reported that isoeugenol induces cell membrane permeability in a reversible and non-disruptive manner. In the present study, TEM image analysis was performed to demonstrate the disruption of the cell membrane (Fig. 3). In the control sample, the cytoplasm, membrane, and cell wall of S. aureus were distinct and clearly visible. However, S. aureus treated with QP-EO exhibited distortions in cell shape and cell wall breakage. In addition, the cell was surrounded by an electron densed-essential oil structure, these aggregated materials were distributed both inside and outside the cell with indistinct cell membranes and cell walls (Fig. 3C). Sotelo-Boyás et al. (2019) observed aggregation around the cells and loss of cell wall in Pectobacterium carotovorum after thyme-loaded nanoparticle treatment, resulting in the leakage of intracellular components. The cell wall of K. pneumoniae treated with Origanum glandulosum and Lavandula dentata L. was surrounded with small particles (Benbrahim et al., 2021). Thus, small particles of essential oil may damage the cell envelope; however, definitive evidence regarding this hypothesis requires further research.

Fig. 3.

Fig. 3

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TEM images of S. aureus. (A) control; (B), (C) and (D) treated with 1.25 mg/mL Q. phillyraeoides EO for 1 h

Biofilm removal efficacy of QP-EO

The inhibitory effect of QP-EO on the initial formation of biofilm is depicted in Figs. 4, 5. When treated at 1/4 MIC and 1/2 MIC for 48 h, a significant (p < 0.05) reduction in total biomass was observed: S. aureus ATCC 25923 showed reductions of 13.0% and 44.9%, S. aureus ATCC 27213 showed reductions of 13.5% and 31.5%, and S. aureus ATCC 29213 showed reductions of 3.8% and 33.6%, respectively. For proteins and polysaccharides, which are key components of the extracellular polymeric substances (EPS), a significant (p < 0.05) reduction was observed when treated at 1/2 MIC and 1/4 MIC for 48 h. In particular, a significantly (p < 0.05) higher protein inhibition capability at 1/2 MIC compared to 1/4 MIC was observed. Considering that microbial growth was gradually inhibited as the QP-EO concentration approached the MIC and excretion of protein strongly related in growth (Ebner et al., 2015), it could be inferred that the lower protein content in S. aureus biofilm at 1/2 MIC was due to the slower microbial metabolism induced by QP-EO. Several previous studies have evaluated the impact of essential oils on biofilm formation inhibition. Abdullah et al. (2021) evaluated the anti-biofilm activity of Elletaria cardamomum essential oil against Escherichia coli O157 and Salmonella Typhimurium JGS 1748. They found significant (p < 0.05) reductions ranging from a minimum of 5% to a maximum of 66.98% at sub-MIC concentrations. Kang et al. (2019) observed the anti-biofilm activity of peppermint essential oil against S. aureus. The formation of S. aureus biofilms decreased with increasing concentrations of peppermint essential oil, observed through environmental scanning electron microscopy analysis. Untreated biofilms were densely structured and encapsulated in EPS, whereas peppermint essential oil-treated biofilms showed significantly disrupted structures, suggesting that essential oil disrupts biofilm structure and reduces biofilm formation.

Fig. 4.

Fig. 4

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Effect of Q. phillyraeoides EO on the biofilm formation of S. aureus ATCC 25923 (A), 27,213 (B), 29,213 (C) determined by crystal violet assay. Different uppercase letters indicate significant (p < 0.05) differences

Fig. 5.

Fig. 5

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Effect of Q. phillyraeoides EO on the biofilm formation of S. aureus ATCC 25923 (A), 27,213 (B), 29,213 (C) determined by calcofluor white staining and FITC staining. Different uppercase letters indicate significant (p < 0.05) differences

QP-EO was also effective against biofilms that had already been established. As shown in Fig. 6, total biomass of S. aureus biofilms treated with QP-EO at 1/2 MIC, MIC, and 2 MIC for 6 h reduced by 20.1%, 32%, and 40.3% for S. aureus ATCC 25923, 18.6%, 34%, and 38.1% for S. aureus ATCC 27213, and 16.8%, 26.7%, and 41.5% for S. aureus ATCC 29213, respectively. For 12 h treatments, the reductions were 24%, 47.7%, and 58% for S. aureus ATCC 25923, 16.1%, 36%, and 50.2% for S. aureus ATCC 27213, and 32.9%, 45.5%, and 47.9% for S. aureus ATCC 29213, respectively. There were no significant (p > 0.05) differences between the 6 h and 12 h treatment groups, suggesting that time may not be a critical factor in reducing biofilm formation and concentration seems to have a greater impact. However, unlike the initial biofilm formation inhibition, there were no significant (p > 0.05) reductions in proteins and polysaccharides compared to the control, suggesting that QP-EO may be more effective in inhibiting the initial formation of biofilms rather than controlling established biofilms.

Fig. 6.

Fig. 6

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Effect of Q. phillyraeoides EO on the preformed biofilm of S. aureus ATCC 25923 (A), 27,213 (B), 29,213 (C) determined by crystal violet assays. Different uppercase letters indicate significant (p < 0.05) differences

Several studies have evaluated the effects of essential oil for biofilm inactivation, but the efficacy differ depending on the type of essential oil. Guo et al. (2019) observed the anti-biofilm activity of Huyou essential oil against Listeria monocytogenes and an increasing trend in biofilm reduction with longer exposure times. They observed thinner 3D structure of biofilm after Huyou essential oil treatment for 8 and 16 h, gradually reduced in thickness, with only a thin monolayer observed after 24 h. Espina et al. (2017) reported that while inactivating biofilms of S. aureus, Escherichia coli, or Listeria monocytogenes is limited with the MIC of carvacrol and citral, treatment with 2 MIC or 4 MIC can effectively control biofilms. These findings suggest that increasing the concentration of QP-EO could reduce EPS production, potentially weakening the resistance of S. aureus biofilms.

In conclusion, this study investigated the antimicrobial activity of QP-EO against S. aureus and presented novel findings that demonstrated significant inhibitory effects with potential bactericidal action. Growth curve analysis indicated delayed bacterial growth, suggesting the potential role of QP-EO in preventing staphylococcal food poisoning. Time-kill kinetics revealed a prolonged tailing effect, indicating irreversible damage to S. aureus following QP-EO treatment. Furthermore, mechanisms underlying the antimicrobial activity of QP-EO include the disruption of the bacterial cell membrane and leakage of cell components. QP-EO also significantly inhibited the initial formation of biofilms and demonstrated efficacy in reducing biomass of established biofilms, although the latter effect was less pronounced. The observed biofilm reduction was concentration-dependent, with higher concentrations of QP-EO leading to greater reductions in total biomass. Despite these promising results, this study has some limitations. This study exclusively focused on S. aureus, and the active compounds and mechanisms underlying the prolonged tailing effect were not explored. Nonetheless, this study provided valuable insights into the antimicrobial potential of QP-EO and its possible applications in microbial control and food safety.

Supplementary Information

Below is the link to the electronic supplementary material.

Supplementary file1 (DOCX 1584 KB) (1.5MB, docx)

Acknowledgements

This work was supported by the research grant of Kongju National University in 2025.

Funding

This study was supported by Kongju National University.

Data availability

Data will be made available on request.

Declarations

Conflict of interest

The authors declare no conflict of interests.

Footnotes

Publisher's Note

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Ye-Seul Yun and Soo-Hwan Kim contributed as co first authors.

References

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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 file1 (DOCX 1584 KB) (1.5MB, docx)

Data Availability Statement

Data will be made available on request.

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