Juniper (Juniperus communis) essential oil was obtained from a commercial supplier located in Antalya, Türkiye. The oil was produced by steam distillation of juniper berries and stored in amber-colored glass bottles at 4 °C until further analysis.

Nanoemulsions were prepared by modifying the method proposed by Joe et al. (2012). Different proportions of juniper EO (2, 4 and 6%), distilled water and 4% Tween 80 (Sigma Aldrich, Taufkirchen, Germany) were used for nanoemulsion formulation. In all nanoemulsion formulations, the concentration of Tween 80 was kept constant at 4%, while the essential oil concentration (2, 4, and 6%) was adjusted by proportionally modifying the aqueous phase.

The components were mixed under appropriate conditions to obtain a homogeneous pre-emulsion. The aqueous phase containing the surfactant was incubated at 86 °C for 1 h to facilitate homogenization and promote nanoemulsion formation, rather than to thermally treat the essential oil. The formulation was maintained under controlled, closed conditions to minimize potential volatilization losses. Gas chromatography–mass spectrometry (GC–MS) analysis was performed on the pure juniper essential oil prior to the nanoemulsion preparation process. After the incubation period, the mixture was cooled to room temperature and pre-sterilized distilled water was added to the cooled mixture at the determined ratios. Homogenization of the nanoemulsions was performed using an ultrasonic probe homogenizer (CY-500 sonicator, Optic Ivymen System, Barcelona, Spain). The homogenization process was applied for 15 min at 95% amplitude. Ultrasonication was performed in continuous mode without pulse application, and the samples were kept in an ice bath during processing while the temperature was continuously monitored to prevent excessive heating. The ultrasonic device used operates at a power of 500 W and a frequency of 20 kHz and includes a ¼ inch titanium alloy probe with a piezoelectric transducer (diameter: 5.6 mm; height: 60 mm). This process was applied in order to reduce the droplet size of the nanoemulsions and ensure a stable structure.

Physicochemical properties of the nanoemulsions were analysed at the METU Central Laboratory (Middle East Technical University, Ankara, Turkey). The droplet size of the emulsions was determined using a Mastersizer 2000 (Malvern Instruments, Malvern, UK) based on laser diffraction principles. The polydispersity index (PDI) was measured by dynamic light scattering (DLS) using a Zetasizer Nano-ZS (Malvern Instruments, Malvern, UK), following the methodology described by Hosseinnia et al. (2017).

In the study, Gram positive Staphylococcus aureus (ATCC29213) and Gram negative Salmonella Paratyphi A (NCTC13) were used as food-borne pathogenic bacteria. Photobacterium damselae, Proteus mirabilis and Vibrio vulnificus were also used as spoilage bacteria. These bacteria were isolated from spoiled anchovy, mackerel and sardine meat (Kuley et al., 2017). These bacteria were selected to test the antimicrobial activity of pure juniper EO and its nanoemulsions, and the study objectives were achieved by analysing fish spoilage and foodborne pathogenic microorganisms.

Antimicrobial activity analysis was carried out to evaluate the effects of juniper EO and nanoemulsions prepared using this oil on foodborne pathogenic and fish spoilage bacteria. These analyses were performed using different methods and aimed to verify and comparatively evaluate the antibacterial activity of nanoemulsions. Within the scope of the analysis, standard methods such as Agar Well Diffusion Assay, MIC and MBC were applied.

In this study, the antimicrobial activity of juniper EO and its nanoemulsions was determined using Mueller-Hinton Agar (MHA, Merck 1.05437, Darmstadt, Germany) according to the well diffusion method of Hwanhlem et al. (2017). Test bacteria were grown in Nutrient broth (Merck 1.05443, Darmstadt, Germany) at 37 °C for 24 h and standardised to 0.5 McFarland cell density (108). Each bacterial cell culture (100 μL) was inoculated into a petri dish containing 20 mL of Muller-Hinton agar. Five 5 mm wells were made in solid medium. Four wells were inoculated with 50 μL of juniper EO in pure or nanoemulsion form. Distilled water was transferred to the other well as control. The negative control was Tween 80. Petri dishes were then incubated at 37 °C for 24 h. After incubation, the zones of inhibition formed around each well were measured in mm using callipers.

MIC and MBC values of pure juniper EO and its nanoemulsions were determined using the CLSI (Clinical and Laboratory Standards Institute, 2006) microdilution method. MIC is defined as the lowest concentration at which the tested substance completely stops the growth of the microorganism, while MBC is the lowest concentration that completely kills the microorganism. The bacterial cultures tested for the experiments were prepared according to the 0.5 McFarland standard and studied using Mueller-Hinton Broth (MHB) medium. For antimicrobial tests, 50 mg/mL stock solutions of juniper EO and nanoemulsions were prepared and diluted to 0.19 μg/mL by the serial dilution method. To each dilution, 1 mL of bacterial suspension was added, and the prepared tubes were incubated at 37 °C for 24 h. After incubation, the turbidity in the tubes was observed, and the lowest concentration without growth was recorded as the MIC value. Following the determination of the MIC values, liquid cultures from the tubes with no growth were inoculated onto Mueller-Hinton agar and incubated at 37 °C for another 24 h. As a result of this second incubation, the lowest concentration at which no bacterial growth was observed was recorded as the MBC value. The reported MIC and MBC values represent the final concentrations of juniper essential oil and nanoemulsion formulations in the test system after dilution and inoculation.

Statistical analyses were performed using one-way analysis of variance (ANOVA) followed by Duncan’s Multiple Comparison Test (SPSS 22, Chicago, Illinois, USA) for post hoc mean separation among treatment groups. All experiments were conducted in triplicate as independent experimental repeats, and data are presented as mean ± standard deviation. Differences were considered statistically significant at p < 0.05.

The average droplet size, polydispersity index (PDI) and surface tension values of the nanoemulsions are presented in Table 1. In the JNEO group with 2% EO content, the relatively low oil phase content resulted in more effective surfactant coverage during the droplet formation process and thus the smallest average droplet size of 21.89 nm was reached. When the essential oil content was increased to 4%JNEO, droplet deposition was promoted due to the increase in viscosity and change in the surfactant/oil phase ratio, resulting in the largest average droplet size of 377.9 nm. PDI values ranged from 0.006 to 0.420; higher PDI values indicated a decrease in the homogeneity of the droplet distribution. The higher PDI value indicates a broader droplet size distribution, suggesting increased polydispersity related to formulation composition. The surface tension measurements decreased to 41.38 mN/m in the JNEO group containing 6% EO, which is attributed to the fact that with the increase in the amount of oil phase, the surfactants form a more organized layer at the water/oil interface, thus reducing the surface tension. The highest surface tension value was 44.24 mN/m in the 2%JNEO group due to the interface instability caused by the low oil phase content. The results obtained in the present study both parallel and differ from the nanoemulsion properties reported in the literature. Joe et al. (2012) reported that the droplet sizes of nanoemulsions prepared from commercial oils such as sunflower seed, castor oil, coconut, peanut and sesame oil ranged between 72.52–875.22 nm; this wide range is thought to be due to the differences in formulation parameters, surfactant types and ratios used in the study. Similarly, some studies have reported nanoemulsions and nanoemulsion-based systems with droplet sizes extending up to approximately 1,000 nm, depending on the formulation composition, oil concentration, and preparation conditions (Costa et al., 2020; Gul et al., 2022; Wilson et al., 2022). Accordingly, the formulations obtained in this study can be defined as nanoemulsion-based systems exhibiting nano-scale properties. Yazgan (2013) reported that sunflower seed oil-based nanoemulsion reached an average droplet size of 381.99 nm; the similarity here is due to the fact that the amount of oil phase directly affects the droplet size. Özogul et al. (2017) reported that nanoemulsions obtained from thyme, rosemary, laurel and sage essential oils formed droplets of 112.82, 63.02, 66.02 and 59.48 nm in size, respectively; Durmus (2020) reported 47.40–94.66 nm droplet size and 35.15–39.18 mN/m surface tension values in formulations based on citrus essential oils (orange, mandarin, grapefruit, lemon). These differences are due to variations in the types of oils used, surfactant selection and nanoemulsion preparation methods.

The GC–MS results presented in Table 2 formed the basis for explaining the chemical composition of juniper essential oil and the potential antimicrobial effects of these components. As a result of the analyses, a total of 30 different compounds were identified, of which the main component was α-Pinene with 90.05%. The high α-Pinene content may have increased the effectiveness of the oil by damaging the microbial cell membrane, as this compound is known to show strong antibacterial and antifungal activity in the literature (Falcão et al., 2018; Najar et al., 2020). Indeed, the presence of other monoterpenes such as α-Myrcene (1.80%) and 2-α-Pinene (1.37%) may have synergistically interacted with this main component to broaden the antimicrobial spectrum.

The α-Cedrol (1.23%) and dl-Limonene (1.18%) contribute to keep juniper oil active against both bacterial and fungal pathogens. In particular, the known antimicrobial effect of α-Cedrol has been enhanced by the presence of this compound. On the other hand, compounds such as Thujone, α-Pinene oxide and Verbenol, although present in the range of 0.01–0.03%, can contribute to the overall antimicrobial activity by altering microorganism cell wall permeability even at low concentrations.

Although the obtained α-Pinene content of 90.05% is in line with the α-Pinene content of 77.11% previously reported in Juniperus sabina L. cones (Ertaş et al., 2019), it showed a higher value depending on the study species and regional ecological conditions. For example, in Juniperus oxycedrus L. subsp. oxycedrus, α-Pinene was found to be 32.85% and β-Myrcene 57.14%, while α-Pinene 19.7% and epi-Sedrol 17.8% were reported in oil obtained from Juniperus excelsa leaves (Gülsoy and Çıvğa, 2016; Doğan et al., 2016). These differences indicate that genetic diversity and geographical conditions among juniper species directly affect essential oil composition.

In conclusion, the component distribution in Table 2 shows that the high content of α-Pinene in juniper essential oil as well as the low concentrations of various monoterpenes and sesquiterpenes support the antimicrobial activity of the oil both directly and through synergistic mechanisms. This facilitated the interaction of the oil with microbial membranes in nanoemulsion formulations, leading to a strong inhibitory effect even at low concentrations.

Inhibition zone diameters measured using the agar well diffusion method (Table 3) clearly demonstrate the antimicrobial activity of juniper essential oil (EO) and its nanoemulsion formulations (JNEO) against the tested bacterial strains. In the group where EO was directly applied, the highest inhibition zone diameters were recorded, indicating more effective disruption of bacterial membrane integrity (p < 0.05). Since Tween 80 lacks antimicrobial qualities, it did not exhibit any inhibition zone.

As shown in Figure 1, inhibition zones increased in a concentration-dependent manner across all tested bacteria, particularly in Vibrio vulnificus and Photobacterium damselae. For V. vulnificus, the inhibition zone diameter increased from 52.25 ± 1.26 mm at 2% JNEO to 64.00 ± 3.46 mm at 6% JNEO, while for P. damselae, zone diameters increased from 15.50 ± 0.58 mm to 35.75 ± 1.89 mm, respectively.

In Staphylococcus aureus and Salmonella Paratyphi A, weak inhibition effects were observed at 2 and 4% JNEO concentrations, whereas at 6%, zone diameters reached 17.00 ± 0.82 mm and 9.25 ± 0.50 mm, respectively. These increases are attributed to the higher concentration of active monoterpenes in EO, which enhances their interaction with bacterial cell wall components. Proteus mirabilis showed high susceptibility to the pure essential oil, while its response to the nanoemulsion formulations was comparatively lower (Table 3).

Similar findings were reported by Taşar et al. (2022), who observed inhibition zones ranging from 8.13 ± 0.67 to 13.43 ± 0.31 mm for Staphylococcus aureus using Juniperus excelsa and J. foetidissima leaf oils, and by Aslan et al. (2021), who reported inhibition zones up to ≥50 mm depending on oil composition and bacterial strain.

The inhibitory effect varied depending on bacterial species and EO concentration. At higher concentrations, JNEO exhibited pronounced antibacterial effects against V. vulnificus and P. damselae, while P. mirabilis showed high susceptibility to the pure essential oil but comparatively lower responses to the nanoemulsion formulations.

The antibacterial efficacy of nanoemulsion formulations was frequently less potent than that of pure EO, despite improvements in transport and stability. Furthermore, in line with previous research, the inhibitory effect of nanoemulsions on bacteria grew as the EO concentration rose (Kayiran et al., 2025). In this work, 2, 4%, or 6% of the EO were incorporated into EO-based nanoemulsions. As a result, their levels of bioactive EO molecules are decreased. This could account for the higher antibacterial activity of pure EO compared to the nanoemulsion form (Özogul et al., 2021). Increased inhibitory zone is seen when the data are recalculated to account for the actual amount of pure EO (Zahi et al., 2017). Therefore, considering the EO content of the nanoemulsions, the antimicrobial efficiency of the nanoemulsion system increased with the addition of EO. These findings suggest that juniper EO and their nanoemusions forms can be considered a promising natural antimicrobial agent, particularly against Gram-negative marine bacteria, and that concentration control plays a critical role in optimizing formulation efficacy.

MIC and MBC values of Juniper EO and nanoemulsions on five different bacterial strains tested are given in Table 4. Tween 80 showed no bactericidal or inhibitory effects on any examined bacteria. MIC and MBC values of Juniper EO differ according to bacterial species. Against Staphylococcus aureus, the MIC and MBC of juniper EO was 0.78 6.25 mg/ mL, whilst towards S. paratyphi A, it exerted the respective MIC and MBC of 12.5 and 25 mg/ mL Juniper EO exhibited similar MIC and MBC against Vibrio vulnificus and Photobacterium damselae, with respective value of 1.56 and 12.5 mg/mL, indicating that the effectiveness of juniper EO on these species was high (Figure 2).

In, the MIC value was 1.56 mg/mL and the MBC value was 12.5 mg/mL and a completely bactericidal effect was observed as the concentration increased. In Proteus mirabilis, the MIC value was 12.5 mg/mL and MBC value was >100 mg/mL and it is understood that this species shows comparatively lower susceptibility to the nanoemulsion formulations, despite its clear sensitivity to the pure juniper EO. As a result of MIC and MBC analyzes, the following findings stand out:

S. aureus and V. vulnificus species are the most susceptible bacteria to juniper EO. Effective inhibition was achieved with low MIC values (0.78–1.56 mg/mL). Similarly, effective inhibition was achieved at low concentrations in P. damselae species, but higher concentrations are required for bactericidal effect. Inhibition and bactericidal effect were observed at higher concentrations in Salmonella Paratyphi A and P. mirabilis species. Especially in Proteus mirabilis, the MBC value exceeded 100 mg/mL, indicating reduced bactericidal activity despite the clear inhibitory effect observed with the pure essential oil.

There are various studies in the literature on the antimicrobial activity of juniper EO. Yazgan (2013) conducted a study on the antimicrobial activity of sage EO and its nanoemulsion form on bacteria causing fish spoilage (P. mirabilis, P. damselae, V. vulnificus, Enterococcus faecalis, Pseudomonas luteola and Serratia liquefaciens). MIC and MBC values were analyzed. In the study, P. luteola and S. liquefaciens species were found to be more sensitive to sage EO compared to other bacteria. Sage EO and its nanoemulsion form showed a high antimicrobial effect on P. luteola with a MIC value of 6.25 mg/mL. Similarly, these components showed a strong inhibitory effect on Proteus mirabilis with a MIC value of 12.50 mg/mL. However, when the MBC values were analyzed (> 25 mg/mL), it was found that sage EO and nanoemulsion form did not produce a significant bactericidal effect on Proteus mirabilis. These results suggest that sage EO has high inhibitory effects, but its bactericidal effects may be more limited. Özoğul et al. (2020) found that grapefruit peel EO showed bacteriostatic properties against most of the bacteria tested in their analysis by disk diffusion, MIC and MBC methods. However, they reported that only the growth of Salmonella paratyphi A, V. vulnificus and S. liquefaciens was inhibited at EO concentrations up to 25 mg/mL. In addition, Rezaei et al. (2016) reported that coriander EO exhibited antimicrobial activity in the range of 2.5–320 μg/mL. Lee et al. (2011) determined the MIC values of C. indicum EO in the range of 1.25–10 μg/mL. They also observed that this oil effectively inhibited the growth of bacteria such as Bacillus cereus, S. aureus, A. hydrophilia, S. choleraesuis, S. enterica, S. sonnei, V. parahaemolyticus and V. vulnificus. They reported the order of MBC as A. hydrophilia < B. cereus, B. subtilis, V. parahaemolyticus < S. aureus, S. sonnei, V. vulnificus.

These literature data support that juniper EO shows antimicrobial activity against various bacteria. However, antimicrobial activity may vary depending on the type of bacteria, the type of juniper used and the concentration of application. Studies show that juniper oil is effective against both Gram-positive and Gram-negative bacteria and that these properties are due to terpenic compounds. However, Hammer et al. (1999) reported that essential oils are more effective against Gram-positive bacteria, while Gram-negative bacteria are more resistant due to the cell wall structure. In this study, a similar result was obtained and S. aureus was the most sensitive species, while gram-negative P. mirabilis was the most resistant species. This may be explained by the fact that juniper EO can target the cell wall of gram-positive bacteria more easily with its chemical components. Juniper EO, which is reported to be particularly effective against fish spoilage bacteria such as Pseudomonas spp., Shewanella spp. and Aeromonas spp. has the potential to extend the shelf life of fresh fish products. Juniper EO nanoemulsions can act on these bacteria by both breaking down the cell wall and disrupting protein and lipid structures. Studies have shown that juniper EO in nanoemulsion form shows high antimicrobial activity even at lower concentrations compared to its free form. This is attributed to the small size and homogeneous distribution of nanoemulsions, as it increases the capacity of the oil to penetrate bacterial cells (Weiss et al., 2009).

In the literature, there are studies examining the effect of essential oils such as thyme, lemon, grapefruit and laurel on pathogenic bacteria. Bhargava et al. (2015) reported the effect of thyme oil nanoemulsions on Listeria monocytogenes, Salmonella Typhimurium and Escherichia coli O157: H7 and observed a reduction of 3.57, 3.26 and 3.35 log CFU/g at 0.1% concentration, respectively. Between Bhargava et al. (2015) and this study, it is noteworthy that inhibition zone diameters were higher in gram-positive bacteria. However, in this study, it was observed that juniper oil provided similar activity even at lower concentrations. Özoğul et al. (2020) reported MIC values as 1.25 mg/mL and 0.75 mg/mL in studies conducted with nanoemulsions containing thyme oil against Salmonella Paratyphi A and V. vulnificus bacteria, respectively. In this study, juniper EO showed a high activity with a lower MIC value of 0.5 mg/mL for V. vulnificus. These differences are thought to be due to the chemical components of the essential oils used. The components of juniper oil with high antimicrobial properties such as α-pinene, sabinene and terpinen-4-ol may explain these differences in efficacy.

The results indicate that juniper essential oil (EO) exhibits strong antimicrobial activity against both Gram-positive and Gram-negative bacteria, particularly when applied in nanoemulsion form. This finding is largely consistent with previous reports on the antibacterial efficacy of essential oils used as natural preservatives. The low MIC and MBC values, together with the wide inhibition zones, clearly demonstrate the potent antimicrobial effect of juniper oil and its nanoemulsion formulations.

The antibacterial activity of juniper nanoemulsions increased consistently in a concentration-dependent manner across all tested species. Although the nanoemulsion formulations exhibited a clear dose-dependent inhibitory effect, pure juniper essential oil showed lower MIC and MBC values under the tested in vitro conditions. Collectively, these results emphasize the strong potential of JNEO as an effective natural antimicrobial agent, particularly against Gram-negative marine bacteria associated with fish spoilage.