A total of 120 samples including raw milk and Kareish cheese samples (60 of each) were collected from diverse sites across Egypt, including Mashtoul Elsouk and Minya Alkamh (Sharkia Governorate), Shibin El-Qanater (Qalyubia Governorate), and El-Bagour (Menoufia Governorate). The samples were collected using aseptic techniques, promptly transported under refrigeration to the laboratory, for immediate bacteriological examination.

The isolation and detection of Listeria species were performed in accordance with the ISO11290-1 standard method (ISO, 1996). The samples underwent a pre-enrichment process utilizing half Fraser broth (Oxoid, Basingstoke, UK) for a duration of 48 h at 37°C, followed by a 24-h enrichment period in Fraser broth (Oxoid, Basingstoke, UK) at the same temperature. Following the enrichment phase, the cultures were transferred onto Palcam agar and Oxford agar (Oxoid, Basingstoke, UK) and incubated at 37°C for a duration of 24 to 48 h. Colonies suspected to be Listeria, exhibiting a black coloration with a surrounding black zone due to esculin hydrolysis, were further characterized through Gram staining and a series of biochemical tests, including oxidase, catalase, CAMP, and motility tests (Gasanov et al., 2005).

Genomic DNA was extracted from bacterial isolates using the QIAamp DNA Mini Kit (Catalogue no. 51304) in accordance with the manufacturer’s guidelines. The extracted DNA’s quality and concentration were evaluated with a NanoDrop spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). The DNA was subsequently used for identification of Listeria species via PCR amplification and agarose gel electrophoresis. For the detection of L. monocytogenes, species-specific primers targeting the 16S rRNA gene (Forward: 5′-GGA CCGGGGCTA ATA CCG AAT GAT AA-3′; Reverse: 5′-TTC ATG TAG GCG AGT TGC AGC CTA-3′) were used, amplifying a 1,200 bp product (Wiedmann et al., 1993). Additionally, primers specific to other Listeria species including L. innocua (Forward: 5′-GGAGTTATTAACGAAGATACT-3′; Reverse: 5′-TTCTGCTTTTACTTCTTTAGCA-3′, L. ivanovii (Forward: 5′-AAGCTGCAGTTATTCATTCC-3′; Reverse: 5′-ATCTAAGAATTTTTGTTTTAGT-3′), L. welshimeri (Forward: 5′-TTCTCGTATTATCGGTTTACCA-3′; Reverse: 5′-GCTTCAAGATAGATTTCTTTCAA-3′, and L. marthii (Forward: 5′-AGAATATATTTGGAACAGCATC-3′; Reverse: 5′-GTTCGATTGCACGGATGGAAAG-3′ were used (Kim et al., 2015). The amplification reaction comprised 12.5 μl of Emerald Amp GT PCR Master Mix (Takara, Code No. RR310A), 1 μl of each forward and reverse primers (20 pmol), 6 μl of template DNA, and nuclease-free water to a final volume of 25 μl. The PCR was performed using a Thermal Cycler (Biometra T3 Thermocycler, Göttingen, Germany). The cycling conditions for L. monocytogenes included an initial denaturation at 94°C for 3 min, followed by 35 cycles of denaturation at 94°C for 1 min, annealing at 60°C for 2 min, and extension at 72°C for 1.5 min. The annealing temperature for other Listeria species was set to 54°C for a duration of 1 min. The PCR products, accompanied by suitable positive and negative controls for each run, underwent analysis through 1% agarose gel electrophoresis. The gel underwent staining with ethidium bromide (Sigma-Aldrich, Germany) and was subsequently visualized under UV light utilizing a gel documentation system (Alpha Innotech, CA, USA).

The antibiotic susceptibility of the recovered isolates was assessed using the Kirby-Bauer disk diffusion method (Bauer et al., 1966). The isolates were tested with a set of 22 antibiotic discs (Oxoid, UK) that are frequently employed in the management of microbial infections (Supplementary Table 1). The diameters of the inhibition zones were carefully measured to the nearest millimeter (mm) using a calibrated ruler. The isolates were classified as susceptible (S), intermediate (I), or resistant (R) to each antibiotic based on the standard reference values set by the Clinical and Laboratory Standards Institute (CLSI) (Clinical and Laboratory Standard Institute, 2023) and the European Committee on Antimicrobial Susceptibility Testing (EUCAST) (Testing, 2024). The evaluation of the multiple antibiotic resistances (MAR) index was calculated to determine the extent of antibiotic resistance. The MAR index reflects the proportion of antibiotics that bacteria have developed resistance to in relation to the total number of antibiotics tested (Krumperman, 1983).

The capacity of L. monocytogenes isolates to form biofilms was assessed through the microtiter plate assay (Djordjevic et al., 2002). Briefly, each isolate was cultured overnight in Tryptic Soy Broth (TSB) (Himedia, Mumbai, India) at a temperature of 37°C. A 100 μl aliquot of the bacterial suspension (10⁷ CFU/ml) was placed into the wells of a sterile 96-well polystyrene microplate, followed by incubation at 37°C for 48 h to facilitate biofilm formation. Following the incubation period, the plates were gently washed three times using 150 μl of distilled water to eliminate any non-adherent cells. Following a 45-min period of air drying, the plates underwent a dyeing process for an additional 45 min with 150 μl of 1% crystal violet. The plates were rinsed with distilled water to remove any residual stain. To elute the stain from the biofilms, 200 μl of 95% ethanol was added to each well, and then 100 μl of the destained solution was transferred to a new microtiter plate. The optical density (OD) of the destained solution was measured at 595 nm using a microtiter plate reader (ELx808-BioTek Instruments, Inc., Burlington, VT, USA). The isolates were categorized into non-biofilm formers (NBF), weak biofilm formers (WBF), moderate biofilm formers (MBF), and strong biofilm formers (SBF) based on the OD values (Stepanovic et al., 2000). The classification was performed using the optical density cut-off value (OD_C), defined as the mean OD of the negative control plus three standard deviations (SD). The following criteria were applied: OD ≤ OD_C (NBF), OD_C < OD ≤ 2 × OD_C (WBF), 2 × OD_C < OD ≤ 4 × OD_C (MBF), and OD > 4 × OD_C (SBF).

Bacteriophages specific to L. monocytogenes were isolated from manure samples collected from dairy farms, following established protocols with minor modifications (Carlton et al., 2005; Kropinski et al., 2009). Briefly, 10 g of fresh manure were mixed thoroughly in 90 milliliters of sterile saline solution and shaken vigorously for a duration of 15 min. The homogenate was subjected to centrifugation at 10,000 × g for 10 min at 4°C to remove particulate matter, and the supernatant was then filtered through a 0.22 μm membrane filter to ensure the elimination of any contaminating bacteria (Carlton et al., 2005).

To enrich the phage, 10 ml of the filtered supernatant were mixed with 10 ml of double-strength TSB and 1 ml (10⁸ CFU/ml) of an overnight culture of L. monocytogenes (KCh5), a pan-drug-resistant (PDR) isolate with strong biofilm formation capacity. The solution was kept at 37°C for 24 h, with gentle agitation at 120 rpm throughout the duration. Notably, L. monocytogenes does not express flagella at 37°C, which may have limited the isolation of flagella-targeting phages (Gründling et al., 2004). After incubation, the culture was centrifuged at 10,000 × g for 10 min. The supernatant was then filtered through a 0.22 μm membrane filter to isolate the phage lysate (Carlton et al., 2005).

The presence of phages in the lysate was confirmed using a spot test (Hyman and Abedon, 2009). Briefly, after the combination of 3 ml of melted soft agar (0.7% agar in TSB) with 100 μl of an overnight culture of L. monocytogenes, the resulting mixture was applied to tryptic soya agar plates that had already solidified (Himedia, Mumbai, India). Once the soft agar had solidified, 10 μl of the phage lysate was carefully applied to the surface of the agar plate. The plates were allowed to dry before being incubated at 37°C for a duration of 24 h. The presence of distinct areas (plaques) at the specified sites indicated the presence of phages (Hyman and Abedon, 2009).

Phage purification was accomplished using the double-layer agar (DLA) technique (Kropinski et al., 2009). Individual plaques were selected with a sterile pipette tip and resuspended in 1 ml of SM buffer (100 mM NaCl, 8 mM MgSO₄, 50 mM Tris–HCl, pH 7.5). The resuspended phages underwent three rounds of plaque purification to guarantee the isolation of a single phage strain (Kropinski et al., 2009).

The propagation of phages necessitated the addition of 100 μl of the purified phage suspension into 10 ml of an overnight culture of L. monocytogenes in TSB. The solution was maintained at 37°C for 6–8 h with gentle agitation at 120 rpm until the bacterial culture was fully lysed. The lysate underwent centrifugation at 10,000 × g for 10 min to eliminate bacterial debris, followed by filtration of the supernatant through a 0.22 μm membrane filter. The phage titer was assessed through the DLA method, and the lysate was preserved at 4°C for short-term applications or at −80°C in SM buffer with 15% glycerol for extended storage (Byun et al., 2022).

Transmission electron microscopy (TEM) was used to morphologically identify L. monocytogenes phage (vB_LmoP_M15). High-titer phage lysates were prepared as described in section 2.6 and exact drops of the lysate were systematically placed onto 200-mesh copper grids that had a carbon-coated collodion membrane. The grids were subjected to negative staining with a 2% (w/v) aqueous uranyl acetate solution, adjusted to a pH of 4.5. Excess stains were removed, and the grids were allowed to air-dry. The samples were analyzed using TEM (Hitachi H600A, Japan) at 80 kV to determine the morphology and size of the phage particles (Ackermann, 2009).

The genomic DNA of vB_LmoP_M15 was extracted using the DNeasy Blood & Tissue Kit (Qiagen, Cat. No. 69581) in accordance with the protocol outlined by Jakočiūnė and Moodley (Jakočiūnė and Moodley, 2018). Briefly, bacterial DNA and RNA were removed by incubating the lysate with DNase I (1 U/μl) and RNase A (10 mg/ml) for 1.5 h at 37°C (Sambrook and Russell, 2001). The phage capsid was digested using proteinase K (20 mg/ml) at 56°C for a period of 1.5 h. DNA purification process was performed in accordance with the kit’s protocol, and the concentration and quality were assessed using a Nanodrop spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). The DNA was quantified by measuring absorbance at 260 nm, while purity was assessed using OD260/280 and OD260/230 ratios. The purified DNA was digested with FastDigest® enzymes (HinfI, HindIII, and HaeIII) at 37°C for 15 min. The DNA fragments were then separated through agarose gel electrophoresis (0.6% agarose in Tris/Borate/EDTA buffer) and visualized under UV light using a gel documentation system. The aim of the restriction digestion was to obtain a preliminary estimation of the genome size of phage vB_LmoP_M15 based on the restriction patterns.

The thermal stability of vB_LmoP_M15 was investigated to evaluate their survival under a range of temperature conditions as previously described (Mahmoud et al., 2018). Phage suspensions with an initial titer of 10⁹ PFU/ml were exposed to temperatures varying from 30 to 100°C in a water bath incubator. The duration of each temperature treatment was consistently maintained for 10 min. After incubation, the samples were promptly cooled using running tap water to ensure the stability of the phage particles. The confirmation of phage viability was achieved through the DLA method.

The pH stability of vB_LmoP_M15 was assessed using a phage lysate with an initial concentration of 9.2 × 10⁹ PFU/ml. The phages were combined in a sequence of tubes filled with SM buffer, which was modified to various pH levels (ranging from pH 2 to 13) through the addition of sodium hydroxide or hydrochloric acid. The tubes were subsequently incubated at 30°C for a duration of 24 h. Following the incubation period, the count of surviving phages was assessed through the plaque assay technique (Jamalludeen et al., 2007).

The sensitivity of vB_LmoP_M15 phage isolates to UV irradiation was evaluated by exposing 10 ml of phage suspension in sterile Petri plates to UV light at two distinct distances (15 cm and 30 cm) from the UV source. A 30-Watt UV lamp functioning at a wavelength of 254 nm was utilized as the UV source. All experimental procedures were conducted in the absence of light to prevent photoreactivation (Lee and Sobsey, 2011). Phage suspensions were subjected to UV irradiation for various time intervals (0, 20, 40, 60, 80, 100, and 120 min). Following exposure, the infectivity of the phages was assessed utilizing the standard plaque assay method (Nuanualsuwan et al., 2008).

The host range (biological activity) of the phage was determined using the spot test, as previously described (Thiemann et al., 1964; Viazis et al., 2011). The isolated vB_LmoP_M15 with the highest titer was subjected to spot testing against several clinical strains that were recovered previously from milk samples, including Pseudomonas aeruginosa, Klebsiella pneumoniae, Staphylococcus aureus, Escherichia coli, L. monocytogenes, and Streptococcus agalctiae. Twenty microliters of the phage suspension (10⁵ PFU/ml) were inoculated on agar plates previously inoculated with each bacterial strain. Bacterial suspensions were adjusted to 10⁸ CFU/ml prior to spotting. The plates underwent incubation at 37°C for a period of 24–48 h to assess the presence of clear plaques. The presence of a clear lytic zone at the designated site indicated the sensitivity to vB_LmoP_M15. For strains that showed lytic zones in the spot test, phage titers were further quantified using the standard double-layer agar plaque assay and expressed as PFU/ml.

The one-step growth curve of vB_LmoP_M15 phage was established using a standard protocol with slight modifications (Hyman and Abedon, 2009). A culture of L. monocytogenes was incubated in TSB at 37°C for a duration of 18 h. The bacterial culture was subsequently exposed to the vB_LmoP_M15 at a multiplicity of infection (MOI) of 0.1. Following a 5-min incubation at 37°C to facilitate phage adsorption to the bacterial cells, phage titration was assessed using the double-layer agar plate method and quantified as PFU/ml. This indicates the preliminary count of phages. Samples were collected at consistent intervals of 5 min over a total period of 70 min. Then, each sample was diluted and then plated onto agar plates to evaluate the number of infective phages at each specified time point. This facilitated the progression of the growth curve. The latency period was determined as the time between phage adsorption and the first observable increase in PFU, indicating the beginning of phage progeny release. The burst size was established by evaluating the number of new phages released from each infected bacterium, necessitating a comparison between the phage count at the peak of the growth curve and the initial phage count. The burst size indicates the mean quantity of phages that are expelled from every infected bacterium. The experiment was conducted on three distinct occasions to guarantee the reliability of the results, and the average values, along with their standard deviations, were documented.

The adsorption rate of vB_LmoP_M15 was assessed using the method outlined by Karumidze et al. (2013). In brief, the filtered lysate of vB_LmoP_M15 was introduced to a log-phase culture of L. monocytogenes, serving as an indicator strain, at a multiplicity of infection of 0.1 PFU/CFU/ml. At 1-min intervals, samples of 1 ml were collected for the initial 5 min and subsequently filtered using membrane filters (0.45 μm, Millipore). The filtrate was analyzed through the DLA technique to ascertain the titer of non-adsorbed phages. The rate constant for adsorption (K) was calculated using the following equation: K = ( 2.3 / ( B × T ) ) × log ( P 0 / P ) , where K is the adsorption rate constant, B is the bacterial host concentration (10⁸ CFU/ml), T is the time (min), P₀ is the unadsorbed phage concentration at the beginning, and P is the unadsorbed phage concentration at the end.

Pasteurized full-fat milk (3% fat) was obtained from a nearby retail store. Prior to inoculation, the milk was tested to ensure the absence of L. monocytogenes and Listeria-specific phages. Both were not detected in the tested samples. The experimental protocol used for the application assay was adapted from a previously established method (Stone et al., 2020). An overnight culture of L. monocytogenes was prepared, subsequently diluted 1:5 in TSB, and incubated for 3 h at 25°C. Bacterial cells were diluted in maximum recovery diluent (Oxoid Ltd., Basingstoke, UK) to achieve an initial concentration of 3.0 log₁₀ CFU/ml in the milk. Calcium chloride (CaCl2, Merck, Germany) was added to the milk at a final concentration of 18.5 mM to enhance phage activity. The vB_LmoP_M15 was propagated and introduced to the test samples at a Multiplicity of Infection (MOI) of 100, while the control samples remained untreated. All samples were incubated at 30°C for 7 days to mimic potential temperature abuse during handling, storage, or retail display. Bacterial viable counts (CFU/ml) were evaluated immediately following the addition of phage (T0) and subsequently at intervals of days 1, 3, 5, and 7. Samples were subjected to serial dilution to enumerate L. monocytogenes and then plated onto Listeria Chromogenic Agar (Himedia, India). The plates were incubated at 37°C for 48 h.

The analysis of the data was conducted utilizing SPSS version 26 (IBM Corp, Armonk, NY, USA). The chi-square test was employed to examine the variations in the prevalence of L. monocytogenes from diverse origins and to evaluate the differences in the AMR patterns of the isolates from various sources. All experimental procedures were conducted in triplicate, and the results were presented as mean ± standard error of the mean (SEM). One-way ANOVA followed by Tukey’s post hoc test was used to analyze the phage characterization results, including the latent period, burst size from the one-step growth curve, pH stability, and thermal stability. An independent sample T-test was employed to assess the antibiofilm and antibacterial effectiveness of L. monocytogenes phage derived from different sources. The p-values were considered statistically significant if they were less than 0.05. Graphs were produced using GraphPad Prism software version 8 (San Diego, CA, USA) and R-software version 4.0.2.¹

Out of the 120 samples analyzed (60 raw milk and 60 Kareish cheese), 24 (20%) were positive for Listeria species using the ISO11290-1 standard method. Raw milk samples showed a significantly higher prevalence (31.7%, 19/60) compared to Kareish cheese samples (8.3%, 5/60) (p = 0.002). Among the positive isolates, L. monocytogenes was the most prevalent species, with 18 isolates (15%) identified, while L. welshimeri was detected in 6 isolates (5%). The prevalence of L. welshimeri was significantly higher in raw milk samples (10%) compared to Kareish cheese samples (0%) (p = 0.027). Colonies suspected to be Listeria appeared black with a surrounding black zone on selective media due to esculin hydrolysis. The isolates demonstrated characteristics of being Gram-positive rods, were catalase-positive, oxidase-negative, motile at 25°C, and showed distinct hemolysis in the CAMP test. Using species-specific primers targeting the 16S rRNA gene, 18 isolates (15%) were identified as L. monocytogenes by a 1,200 bp band, and 6 isolates (5%) as L. welshimeri by a 237 bp band. As revealed in Table 1, the prevalence of Listeria species varied significantly between raw milk and Kareish cheese samples, with raw milk exhibiting a higher percentage of both L. monocytogenes and L. welshimeri isolates. To further understand the factors influencing this distribution, a random forest classification analysis revealed that season and location were the most critical predictors for L. monocytogenes prevalence.

L. monocytogenes showed a high resistance rate of 95.8% to cotrimoxazole, and L. welshimeri isolates were 100% resistant to the same antibiotic. Resistance to ceftriaxone, another cephalosporin, was also prevalent, with 87.5% of L. monocytogenes and 83.3% of L. welshimeri isolates being resistant. In contrast, all Listeria isolates were fully susceptible to vancomycin (VA), teicoplanin (TEC), tigecycline (TGC), levofloxacin (LEV), and ciprofloxacin (CIP), except for the single pan-drug-resistant isolate (KCh5), which showed resistance to all tested antibiotics (Supplementary Table 1). These findings suggest that the mentioned antibiotics remain effective for most Listeria infections. Intermediate resistance was observed for erythromycin (45.8% in L. monocytogenes and 33.3% in L. welshimeri) and rifampicin (20.8% in L. monocytogenes and 16.7% in L. welshimeri), indicating potential challenges in treatment with these agents. There were statistically significant differences in the resistance rates of Listeria species isolates from raw milk and Kareish cheese samples to ampicillin, streptomycin, chloramphenicol, and trimethoprim/sulfamethoxazole (p = 0.018, 0.042, 0.011, and 0.028, respectively). Meanwhile, there were no statistically significant differences (p > 0.05) in the resistance rates of Listeria species isolates from different origins to the other antimicrobials investigated (Supplementary Table 1). Figures 1A,B illustrates the antimicrobial resistance rates of Listeria species isolates from different sources, highlighting the varying resistance patterns depending on the source of isolation. Figure 2 complements these findings by showing the frequency of resistance to various antimicrobial agents among Listeria isolates from different species (A) and sources (B), providing a detailed overview of the resistance profiles.

Seventeen L. monocytogenes isolates (94.4%) exhibited a multidrug-resistant (MDR) phenotype. There were no statistically significant differences (p > 0.05) in the MAR index among isolates. However, one L. monocytogenes (code no. kch5) isolated from Kareish cheese exhibited complete resistance to all 22 antibiotics tested (MAR index = 1.0), highlighting the emergence of a highly resistant L. monocytogenes strain in the food chain. Figure 3 and Supplementary Figure 1 provide further insights through hierarchical clustering and principal component analysis (PCA), respectively, revealing the diversity and polyclonality of Listeria isolates. The hierarchical clustering dendrogram (Figure 3; Supplementary Figures 1, 2) shows the relatedness of Listeria species isolates based on their antimicrobial resistance profiles. This analysis revealed distinct resistance clusters among isolates from different sources and geographical locations. The PDR isolate (KCh5) from Kareish cheese demonstrated resistance to all antimicrobial classes tested. Several MDR isolates, notably RM12 and RM19 from raw milk, exhibited resistance to multiple antimicrobial classes, particularly β-lactams, cephalosporins, and carbapenems. The clustering analysis further indicated geographical associations in resistance patterns, with isolates from identical regions displaying similar resistance profiles.

The biofilm-forming ability of L. monocytogenes isolates obtained from raw milk and kareish cheese samples was quantitatively assessed using a microtiter plate assay. The isolates were classified into three categories based on their OD values at 595 nm: WBF, MBF, and SBF. The analysis revealed that 38.46% (n = 5/13) of raw milk isolates were classified as WBF, 46.15% (n = 6/13) as MBF, and 15.38% (n = 2/13) as SBF. In contrast, 40% (n = 2/5) of the kareish cheese isolates were WBF and 60% (n = 3/5) were MBF (Figures 4A,B). No significant differences were observed in biofilm-forming capacity between isolates derived from raw milk and kareish cheese (p = 0.68).

Using the spot test and DLA technique, a bacteriophage specific to L. monocytogenes, designated vB_LmoP_M15, was successfully isolated from manure samples. The phage produced clear plaques on lawns of L. monocytogenes, indicating its lytic activity (Supplementary Figure 3). A single plaque was purified and propagated, yielding a high-titer phage lysate. The phage was stored at 4°C for short-term use or at −80°C in SM buffer with 15% glycerol for long-term storage.

The morphology of the isolated phage was examined using TEM following staining with uranyl acetate. The electron micrographs (Figure 5) reveal that the phage displays icosahedral heads accompanied by short non-contractile tails, a morphology characteristic of phages within the Podoviridae family. The phage exhibited head dimensions ranging from 50.42 nm to 56.86 nm, with a notably short tail length, aligning with the distinctive characteristics of the Podoviridae family.

The genomic DNA of the vB_LmoP_M15 phage was successfully extracted and analyzed using restriction enzymes. The extracted DNA was visualized on a 0.6% agarose gel, showing a clear band corresponding to the phage genome. Genome size estimation was performed by comparing the undigested DNA band with a high molecular weight marker, suggesting an approximate genome size of 48.5 kb. Restriction fragment length polymorphism (RFLP) analysis was performed using HinfI, HindIII, and HaeIII enzymes. The digestion patterns revealed distinct banding profiles for each enzyme (Supplementary Figure 4).

The thermal stability of vB_LmoP_M15 was assessed at temperatures between 30 and 100°C over a duration of 10 min. The DLA method was employed to assess viability, with the findings presented in Figure 6A. A significant temperature-dependent reduction in phage survival was observed (p < 0.0001). At the initial temperature, the phage titer was 8.96 ± 0.092 log PFU/ml. Minimal reduction occurred at 30°C (8.41 ± 0.064 log PFU/ml), but viability declined progressively with increasing temperature. At 40°C, the titer dropped to 7.78 ± 0.103 log PFU/ml (~20% reduction), and at 50°C, it decreased to 6.61 ± 0.075 log PFU/ml (~50% reduction), indicating moderate thermostability. However, at 60°C, the titer was 0 log PFU/ml, and no viable phages were detected at temperatures ≥70°C, demonstrating complete inactivation. These results highlight the temperature sensitivity of vB_LmoP_M15, with significant stability at moderate temperatures (up to 50°C) and complete inactivation at higher temperatures (≥60°C). The thermostability at 50°C, suggests potential applications in moderate-heat environments, while the sensitivity to higher temperatures limits its use in high-heat conditions.

The stability of vB_LmoP_M15 to pH values ranging from 2 to 13 for 24 h at 30°C was examined. The results revealed significant pH-dependent survival (p < 0.0001) (Figure 6B). The phage exhibited maximum stability at neutral pH (pH 7) and initial pH, with titers of 8.96 ± 0.087 and 8.96 ± 0.092 log PFU/ml, respectively. Moderate stability was observed at pH 4 (6.95 ± 0.087 log PFU/ml) and pH 5 (7.8 ± 0.116 log PFU/ml), while viability decreased at pH 6 (7.2 ± 0.115 log PFU/ml) and pH 8 (6.18 ± 0.104 log PFU/ml). Extreme pH values (pH 2, 3, and 9–13) resulted in complete inactivation, with no detectable plaques. These findings indicate that vB_LmoP_M15 is highly stable at neutral pH, moderately stable under mildly acidic conditions, and sensitive to extreme pH values. The statistical significance (p < 0.0001) underscores the critical role of pH in phage survival, with potential applications in environments with neutral to slightly acidic conditions.

The sensitivity of vB_LmoP_M15 phage to UV irradiation (254 nm) for varying durations (0 to 120 min) was evaluated. As presented in Figure 6C, there are significant time-dependent reductions in phage survival (p < 0.0001). At 0 min, the phage titer was 8.96 ± 0.098 log PFU/ml, indicating high initial viability. After 20 min, the titer remained stable at 8.93 ± 0.075 log PFU/ml, demonstrating short-term UV resistance. However, viability declined significantly at 40 min (7 ± 0.116 log PFU/ml) and further at 60 min (6.65 ± 0.087 log PFU/ml), indicating moderate UV stability. Prolonged exposure beyond 60 min (80, 100, and 120 min) resulted in complete inactivation, with no detectable plaques. These findings highlight the UV sensitivity of vB_LmoP_M15 phage isolates, with significant stability during short-term exposure (up to 20 min) and moderate stability up to 60 min. Complete inactivation occurred after prolonged exposure, limiting the phage’s utility in high-UV environments.

The adsorption kinetics of vB_LmoP_M15 phage to host cells were evaluated over a 5-min period. Statistically significant differences were observed across the tested time intervals (p < 0.0001) (Figure 7A). At the initial time point (0 min), the phage titer was 8.96 ± 0.192 log PFU/ml. Within the first minute, the titer of non-adsorbed phages decreased sharply to 6.48 ± 0.115 log PFU/ml, indicating efficient and rapid attachment to host cells. This trend continued over the subsequent minutes, with titers of 6.43 ± 0.215 log PFU/ml at 2 min, 6.28 ± 0.162 log PFU/ml at 3 min, 6.23 ± 0.133 log PFU/ml at 4 min, and 6.2 ± 0.116 log PFU/ml at 5 min. These findings indicate that the majority of phage adsorption occurred within the first minute, followed by a gradual stabilization in adsorption efficiency. The high adsorption rate observed in this study underscores the efficiency of vB_LmoP_M15 phage in attaching to L. monocytogenes host cells, a critical step for initiating the lytic cycle. The results provide significant understanding of the initial phases of phage-host interactions and underscore the promise of vB_LmoP_M15 phage as a potent biocontrol agent targeting L. monocytogenes.

The one-step growth curve of vB_LmoP_M15 was determined to elucidate its replication kinetics within L. monocytogenes KCh5 host cells. Figure 7B revealed distinct phases of phage infection, including adsorption, eclipse, replication, and release. The initial phage titer was 7 ± 0.192 log PFU/ml (approximately 1 × 10⁷ PFU/ml). During the first 15 min, the titer decreased to 6 ± 0.116 log PFU/ml (approximately 1 × 10⁶ PFU/ml), reflecting the adsorption of phages to host cells and the subsequent eclipse phase, during which no infectious phages are detected. The latent period, characterized as the duration from adsorption to the initiation of progeny release, was 15–20 min. Following the latent period, a sharp increase in phage titer was observed, indicating the onset of the lytic cycle. The titer increases to 7.67 ± 0.098 log PFU/ml at 20 min, 7.2 ± 0.116 log PFU/ml at 25 min, and 7.75 ± 0.144 log PFU/ml at 30 min. The maximum titer reached was 8.54 ± 0.139 log PFU/ml (approximately 3.47 × 10⁸ PFU/ml) at 35 min, indicating the conclusion of the replication cycle and the subsequent release of progeny phages. The burst size, determined by the ratio of the final phage titer to the initial titer, was 172 phages per infected cell. The findings indicate that vB_LmoP_M15 phage shows efficient replication kinetics, characterized by a relatively short latent period and a substantial burst size, underscoring its potential as an effective lytic phage targeting L. monocytogenes.

The phage displayed differential lytic activity across the tested strains (Supplementary Table 2). Notably, vB_LmoP_M15 exhibited strong lytic effects (+++) against L. monocytogenes and S. aureus, with high titers recorded at 9.2 × 10⁸ and 2.2 × 10⁷ PFU/ml, respectively. Moderate lytic activity (++) was observed against certain strains of S. aureus and L. monocytogenes, with titers ranging between 3.3 × 10⁶ and 9 × 10⁶ PFU/ml. In contrast, weak lysis (+) was detected for E. coli and S. agalactiae, yielding titers of 1.5 × 10⁶ and 2.7 × 10⁶ PFU/ml, respectively. No lytic activity was observed against P. aeruginosa. These titers were determined using plaque assay for strains that showed sensitivity in the spot test, while the spot test was used qualitatively to assess initial lytic potential. Collectively, these findings indicate that the isolated phage possesses a relatively broad host range, with marked efficacy against L. monocytogenes and S. aureus.

The vB_LmoP_M15 phage demonstrated a strong antibiofilm activity. The reduction in biofilm biomass was consistent across both preformed biofilms and inhibition of biofilm formation, highlighting the phage’s potential as an effective antibiofilm agent (Supplementary Table 3; Figures 8–10). There were statistically significant differences (p < 0.05) in biofilm formation between all control untreated biofilms and phage-treated biofilms at both the preformed biofilm and inhibition of biofilm formation (Supplementary Table 3). These findings underscore the phage’s ability to inhibit biofilm formation and disrupt established biofilms, making it a promising candidate for controlling L. monocytogenes.

The efficacy of vB_LmoP_M15 for inhibition of L. monocytogenes (KCh5) growth in pasteurized milk was evaluated. Pasteurized milk samples, which were initially inoculated with L. monocytogenes (3.0 log10 CFU/ml), underwent treatment with vB_LmoP_M15 phage at a Multiplicity of Infection (MOI) of 100 and were incubated at 30°C for a duration of 7 days.

In the untreated control samples, L. monocytogenes exhibited substantial growth, with counts increasing from the initial 3.0 log₁₀ CFU/ml to 7.55 log₁₀ CFU/ml by day 7 (Figure 11). Conversely, treatment with vB_LmoP_M15 phage significantly inhibited the growth of L. monocytogenes. While bacterial counts in the phage-treated group increased to 5.1 log₁₀ CFU/ml by day 7, this represented a marked attenuation of growth compared to the control.

A significant reduction in L. monocytogenes counts was observed in the phage-treated milk relative to the untreated control milk starting from day 1. Specifically, a log₁₀ reduction of 0.523 (p = 0.011) was recorded by day 1. This inhibitory effect became more pronounced over the incubation period, with log₁₀ reductions reaching 1.200 (p = 0.001) by day 3, 1.723 (p < 0.0001) by day 5, and culminating in a maximum observed log10 reduction of 2.45 (p < 0.0001) by day 7 (Figure 11).