The frozen strains of E. coli, S. aureus, L. monocytogenes and S. typhimurium were revived by inoculating them on NA solid medium. Single colonies were picked and transferred into NB liquid medium, where they were cultured at 37°C for 24 h (Su et al., 2023). The bacterial cultures were then stored at 4°C. Before starting the experiments, the bacterial suspensions were adjusted to a concentration of 10⁸ CFU/mL using a McFar-land turbidity standard.

The antibacterial activity of MEO against common foodborne bacteria (E. coli, S. aureus, L. monocytogenes, and S. typhimurium) was evaluated using the filter paper disk diffusion method (El Barnossi et al., 2020). A 100 μL bacterial suspension with a concentration of 1 × 10⁶ CFU/mL was spread evenly on NA agar plates using a sterile spreader. Sterile filter paper disks were gently placed at the center of each plate. MEO (5 μL) was added to the filter paper disks, and sterile water was used as a blank control. The plates were incubated at 37°C for 24 h. The inhibition zones were measured and photographed. This procedure was repeated three times for accuracy.

The MIC and MBC of MEO were determined using the microdilution method (Veiga et al., 2019). Initially, a solution of MEO in NB medium was prepared at a concentration of 60 μL/mL, with 4% DMSO added as a cosolvent. This solution was then further diluted to concentrations of 10, 9, 8, 7, 6, 5, 4, 3, 2 and 1 μL/mL. In a sterile 96-well plate, 100 μL of the diluted MEO solution was added to each well from left to right, followed by the addition of 100 μL of bacterial suspension (1 × 10⁶ CFU/mL), gently mixing the contents. The plate was then incubated at 37°C for 24 h in a constant-temperature microbiological incubator. The bacterial growth in each well was observed and analyzed, with the lowest concentration showing no bacterial growth and no increase in optical density (OD) defined as the MIC value (Kowalska-Krochmal and Dudek-Wicher, 2021). The incubation continued for an additional 48 h, and the growth was reassessed. The lowest concentration with no bacterial growth and no increase in OD was defined as the MBC value. The experiment was repeated three times. The blank group did not contain MEO or DMSO, while the control group only included DMSO.

The determination of growth curve referred to the method of Feng et al. (2022). The effect of MEO on the growth curve of the test bacteria was assessed in a sterile 96-well plate. MEO solutions at concentrations of 0.25 MIC, 0.5 MIC, and 1 MIC, determined from preliminary experiments, were added sequentially, with 200 μL of each solution introduced into the wells. NB medium without MEO served as the blank control. Subsequently, 20 μL of bacterial suspension (1 × 10⁶ CFU/mL) was added to each well, and the plate was incubated at 37°C in a shaking water bath at 100 rpm. The OD at 600 nm was measured and recorded every 2 h to plot the growth curve. Each group was tested in triplicate, and the bacterial growth curve was represented by the average turbidity, with absorbance corresponding to the time intervals.

The determination of the contents is based on the method of Liu et al. (2020). In a 10 mL suspension of test bacteria (1 × 10⁶ CFU/mL), MEO was added, using 4% DMSO as a co-solvent, to achieve a final concentration equal to the MIC. The blank group consisted of samples without MEO or DMSO, while the control group included only DMSO. The bacterial suspension was then incubated at 37°C in a shaking water bath at 100 rpm for 2 h. Following incubation, the suspension was centrifuged at 8000 rpm for 10 min to collect the supernatant. This supernatant was transferred to fresh tubes for analysis, where the optical density at 260 and 280 nm was measured to quantify the nucleic acids and proteins released from the cytoplasm.

The solution conductivity was determined according to the method of Wingfield (2014). In a 10 mL suspension of test bacteria (1 × 10⁶ CFU/mL), MEO was added, with 4% DMSO used as a co-solvent, to achieve a final concentration corresponding to the MIC. The blank group consisted of samples without MEO or DMSO, while the control group included DMSO only. The bacterial suspension was incubated at 37°C in a shaking water bath at 100 rpm for 2 h. After incubation, the suspension was centrifuged at 8,000 rpm for 10 min to collect the supernatant, which was then transferred to fresh tubes. The conductivity of each tube’s supernatant was measured using a conductivity meter.

The determination of intracellular enzymes referred to previous research methods (Shi et al., 2016; Rubeena et al., 2020; Li et al., 2019). In a 10 mL suspension of test bacteria (1 × 10⁶ CFU/mL), MEO was added, along with 4% DMSO as a co-solvent, to achieve a final concentration at the MIC. The blank group consisted of samples without MEO or DMSO, while the control group included DMSO only. The bacterial suspension was then incubated at 37°C in a shaking water bath at 100 rpm for 2 h. Following this, the suspension was centrifuged at 8,000 rpm for 10 min, discarding the supernatant and collecting the bacterial pellet. High-efficiency RIPA lysis buffer was added to the cells, and a cell disruptor was used to thoroughly lyse the cells while in an ice bath. The lysate was centrifuged again at 8,000 rpm for 10 min, and the supernatant was transferred to a new tube and kept on ice for subsequent biochemical assays. The detection steps for AKP, ATP, MDA, and SOD activities are detailed in the kit instructions.

To confirm and validate the results observed under an optical microscope, we slightly modified the previously reported method of observing test bacteria using SEM (Bai et al., 2019). The bacteria (at a concentration of 10⁸ CFU/mL) were treated with MEO at MIC concentration for 2 h at 37°C. The precipitate was then centrifuged at 6,000 rpm for 10 min and washed three times with PBS. The collected cells were fixed in 2.5% glutaraldehyde at 4°C for 24 h, followed by dehydration using a series of ethanol concentrations (15%, 30%, 45%, 60%, 75%, 90%, and 100%) for 10 min each. Finally, the dehydrated samples were gold-coated and examined using SEM. A control experiment was conducted without MEO treatment.

To prepare the MEO solutions, anhydrous ethanol was used to create concentration gradients of 1.25, 2.5, 5, 10, 15, 20, and 40 mg/mL. A 5 mg/mL BHT solution served as the positive control. For each test, 0.5 mL of the sample solution was mixed with an equal volume of a 0.2 mM DPPH anhydrous ethanol solution. The mixture was thoroughly mixed and then placed in the dark for 30 min, with three parallel operations for each concentration. The absorbance was measured at a wavelength of 517 nm (Ashmawy et al., 2024). The experiment was repeated three times. The DPPH radical scavenging rate was calculated using the following formula (1):

A₀ represents the absorbance of the sample group (sample without DPPH solution), A₁ represents the absorbance of the sample group containing DPPH, and A₂ rep-resents the absorbance of the control group (sample without DPPH solution).

To prepare the MEO solutions, anhydrous ethanol was used to create concentration gradients of 1.25, 2.5, 5, 10, 15, 20, and 40 mg/mL, with a 5 mg/mL BHT solution serving as the positive control. A 9.0 mmol/L FeSO4 solution, a 9.0 mmol/L ethanol-salicylic acid solution, and a 3% H2O2 solution were prepared and stored after thorough mixing. In a test tube, 1 mL of the sample solution was taken, followed by the addition of 1 mL of the prepared FeSO4 solution and 1 mL of the ethanol-salicylic acid solution. Finally, 1 mL of the H2O2 solution was added. The mixture was shaken thoroughly and incubated in a water bath at 37°C for 30 min. The absorbance was then measured at a wavelength of 510 nm (Zhu et al., 2024). This experiment was repeated three times. The hydroxyl radical scavenging rate was calculated using the following formula (2):

A₀ represents the absorbance value of distilled water participating in the reaction instead of the sample solution. A₁ represents the absorbance value of the sample solution participating in the reaction. A₂ represents the absorbance value of the sample solution and distilled water instead of 3% H2O2 in the reaction.

To prepare the MEO solutions, anhydrous ethanol was used to create concentration gradients of 1.25, 2.5, 5, 10, 15, 20, and 40 mg/mL, with a 5 mg/mL BHT solution serving as the positive control. In a 10 mL centrifuge tube, 1 mL of the solution was centrifuged, and then 2.5 mL of phosphate buffer (0.2 mol/L, pH 6.6) and 2.5 mL of 1% potassium ferricyanide solution were added. The mixture was thoroughly mixed and reacted at 50°C for 20 min before being rapidly cooled. Next, 2.5 mL of 10% tri-chloroacetic acid solution was added, followed by centrifugation at 4,000 rpm for 15 min. The supernatant (2.5 mL) was then taken and mixed with 2.5 mL of distilled water and 0.5 mL of 0.1% ferric chloride solution. After sufficient mixing, the reaction was allowed to proceed for 10 min, and the absorbance was measured at a wave-length of 700 nm. This experiment was repeated three times to ensure accuracy and reliability (Chebbac et al., 2023).

The inhibition ability of MEO was evaluated using the disk diffusion method. As shown in Figure 2, MEO exhibited growth inhibition against E. coli, S. aureus, L. monocytogenes, and S. typhimurium. The diameters of the inhibition zones are presented in Table 2, with the inhibition zones for E. coli, S. aureus, L. monocytogenes, and S. typhimurium measured at 12.63 ± 0.35 mm, 10.94 ± 0.44 mm, 12.63 ± 0.35 mm, and 12.5 ± 0.24 mm, respectively. These results demonstrate that MEO has significant antibacterial potential.

The MIC and MBC values of MEO against four bacterial strains were determined using the broth microdilution method, with the results shown in Table 3. The MIC values for E. coli, S. aureus, L. monocytogenes, and S. typhimurium were 5, 5, 4, and 4 μL/mL, respectively. The corresponding MBC values were 5, 8, 5, and 5 μL/mL, respectively. These results indicate that MEO exhibits relatively weaker bactericidal activity against S. aureus compared to the other strains. In the control group, bacterial growth was normal, confirming that the addition of a certain amount of DMSO had no inhibitory effect on bacterial growth.

The effect of MEO on the growth of E. coli, S. aureus, L. monocytogenes, and S. typhimurium is shown in Figure 3. In the control group, all four bacteria rapidly entered the logarithmic growth phase. However, treatment with 0.25 MIC and 0.5 MIC of MEO effectively delayed the logarithmic growth phase for all test bacteria. At the 12 h mark, different concentrations of MEO exhibited inhibitory effects on the four bacterial strains. The optical density (OD) values at 1 MIC remained almost unchanged within 12 h, indicating that the test bacteria were highly sensitive to this concentration. Significant differences were observed between the MEO-treated groups and the control group.

The release of intracellular substances after MEO treatment was measured using UV spectrophotometry. As shown in Figure 4, after incubating the four bacterial strains with MEO for 2 h, the absorbance at 260 and 280 nm significantly increased. A statistically significant difference (p < 0.05) was observed between the test group and both the blank and control groups. The rise in extracellular nucleic acids and proteins suggests that MEO disrupts the permeability of the bacterial cell membranes, leading to the leakage of intracellular materials and consequently inhibiting bacterial growth and reproduction.

The electric conductivity of bacterial cultures was measured after incubation with MEO using a conductivity meter. As shown in Figure 5, after 2 h of incubation, the conductivity of the test group significantly increased compared to the blank group (p < 0.05), which aligns with the findings in Figure 4. This increase in conductivity is due to the leakage of intracellular substances such as nucleic acids and proteins, which carry charges and elevate the ionic concentration in the culture medium. In contrast, the conductivity of the control group slightly decreased compared to the blank group, likely because DMSO, a non-ionic solvent, does not contribute to ion concentration and may reduce the number of conductive ions per unit volume.

Alkaline phosphatase enzyme is closely related to bacterial metabolic activity, membrane stability, and the synthesis and degradation of the cell wall. When the level of AKP decreases, it indicates damage to the bacterial cell membrane or wall, leading to reduced metabolic activity and slower growth. Therefore, a reduction in AKP enzyme levels can be considered a marker of bacterial damage (Zhou et al., 2022; Shi et al., 2024). As shown in Figure 6A, after 2 h of co-incubation with MEO, the AKP levels in all four test bacteria were significantly reduced. Furthermore, there was a notable difference between the test group and the blank and control groups (p < 0.05). This suggests that MEO has a destructive effect on the bacterial cell wall.

ATPase plays a crucial role in bacterial cells by hydrolyzing ATP into ADP, releasing energy necessary for various cellular processes. When the level of ATPase decreases, the bacteria experience multifaceted damage (Mao et al., 2024). As illustrated in Figure 6B, after 2 h of incubation with MEO, the ATPase content in all four test bacteria showed a significant reduction. Moreover, there was a noticeable difference between the experimental group and the blank and control groups (p = 0.05). A reduction in ATPase levels leads to insufficient energy metabolism, impaired transmembrane ion transport, membrane potential imbalance, and weakened stress responses. In severe cases, these disruptions can inhibit bacterial growth or even lead to cell death.

Malondialdehyde is one of the primary markers of lipid peroxidation in cell membranes, typically produced under oxidative stress conditions. An increase in MDA levels often indicates that the bacteria have undergone oxidative damage or that their membrane structure has been compromised (Suo et al., 2022; Chen et al., 2022). As shown in Figure 7A, after 2 h of co-incubation with MEO, the MDA content in the four test bacteria significantly increased. There was a marked difference between the experimental group and the blank and control groups (p < 0.05). The increase in MDA levels is a sign of severe oxidative damage to bacterial cells, representing structural damage to the membrane and heightened oxidative stress, which negatively impacts bacterial growth and reproduction.

Superoxide dismutase primarily functions to eliminate superoxide anions in bacteria by catalyzing their conversion into hydrogen peroxide and oxygen, thereby protecting cells from oxidative damage. An increase in SOD levels typically indicates that the bacteria are experiencing oxidative stress (Mohammad et al., 2014). As shown in Figure 7B, after 2 h of co-incubation with MEO, the SOD enzyme content significantly increased in the four test bacteria. There were statistically significant differences between the experimental group and the blank and control groups (p < 0.05). The elevated SOD levels reflect the bacterial response to oxidative stress, indicating that the bacteria are under environmental pressure, which may involve membrane damage, oxidative damage to proteins and DNA, and metabolic dysfunction.

The impact of MEO on the cellular morphology of the four test bacteria was observed using SEM. As shown in Figure 8, the control group displayed bacteria with smooth, intact surfaces, regular edges, full bodies, and tightly packed structures. However, after treatment with MEO at the MIC level, varying degrees of deformation were evident in all four bacteria. The bacterial surfaces became unclear, collapsed, with cell membrane indentations, ruptures, and distorted shapes. These results indicate that MEO damaged the bacterial cell wall and membrane structures. The disruption of these structures led to the leakage of intracellular contents, compromising bacterial integrity and resulting in its antibacterial effect.