Surface-dependent persistence and transfer of Listeria monocytogenes in multispecies biofilms on food-contact surfaces

Abstract

Listeria monocytogenes persistence in food processing environments is closely associated with biofilm formation on food-contact surfaces and subsequent transfer to fresh produce. However, the role of native environmental microbiota in shaping pathogen behaviour within biofilms remains poorly understood. This study evaluated biofilm formation, transfer dynamics, cell surface hydrophobicity, and biofilm organization of L. monocytogenes in mono- and multispecies biofilms on stainless-steel (SS), silicone rubber (SR), and polycarbonate (PC) surfaces under low-shear conditions. Multispecies biofilms enhanced L. monocytogenes persistence and modulated cross-contamination dynamics in a surface-dependent manner. On SS, multispecies biofilms showed reduced initial transfer but sustained cumulative transfer across sequential contacts compared to monospecies biofilms, indicating greater structural stability. In contrast, PC exhibited high initial transfer followed by rapid transfer plateauing, whereas SR supported limited accumulation after the first contact event. Higher cell surface hydrophobicity in multispecies biofilms, particularly on SS, was associated with increased biofilm cohesion. Together with fluorescence microscopy observations, these results suggest that biofilm organization, rather than total cell numbers alone, plays a key role in determining persistence and transfer behaviour. Overall, both surface properties and microbial community composition influence L. monocytogenes biofilm stability and cross-contamination risk. Incorporating surface-dependent and ecological factors into hygienic design and sanitation strategies may improve contamination control in fresh produce processing environments.

1. Introduction

Listeria monocytogenes is a major foodborne pathogen of global concern due to the severity of listeriosis and its disproportionate impact on public health. Although the incidence of the disease is relatively low, listeriosis is associated with high hospitalization and mortality rates, particularly among vulnerable populations [1]; [2,3]. Both sporadic cases and large-scale outbreaks of listeriosis have been linked to a wide range of foods, with ready-to-eat products and, increasingly, fresh produce representing major vehicles for Listeria monocytogenes transmission due to the absence of a final lethal step and the consumption of fruits and vegetables in raw form [[4], [5], [6]]. Fresh spinach has been implicated in Listeria-related recalls and outbreaks, highlighting its vulnerability to post-processing contamination and its relevance as a model commodity for cross-contamination studies ([7], 2026; [8]).

The food safety significance of Listeria monocytogenes is further amplified by its ability to persist in natural, food-associated, and processing environments. Food processing facilities are characterized by the extensive use of diverse food contact surfaces that provide favourable conditions for microbial attachment and biofilm development, a key factor contributing to the long-term persistence of L. monocytogenes in these settings [[9], [10], [11], [12]]. The physicochemical properties of food contact surfaces, including roughness, hydrophobicity, surface charge, and conditioning by organic residues, play a critical role in determining attachment efficiency and biofilm development [13]; [11]. Stainless steel, widely used in the fresh produce and food processing industries due to its durability and cleanability, can nonetheless support bacterial attachment and biofilm formation, particularly when surface imperfections or organic conditioning layers are present [14]; [15].

Most studies investigating L. monocytogenes biofilms have traditionally relied on monospecies models, which have provided valuable insights. However, these simplified systems do not fully capture the complexity of food processing environments, where surfaces are colonized by diverse microbial communities. In recent years, increasing attention has been directed toward dual- and multispecies biofilms, which better reflect the ecological interactions occurring in situ and may significantly influence pathogen behaviour, including persistence and stress tolerance [16,17]. Despite this shift, most multispecies biofilm studies employ defined consortia composed of a limited number of well-characterized strains, which may not adequately represent the diversity and variability of resident microbiota in processing facilities. While recent studies have started to characterize biofilms formed by native environmental microbiota in fresh-cut produce processing environments [9], their functional role in shaping L. monocytogenes behaviour and transfer dynamics remains largely unexplored. These interactions may modulate L. monocytogenes behaviour within biofilms, but their impact on pathogen transfer under fresh produce processing conditions remains poorly understood.

Biofilm-associated characteristics such as structural organization, cell surface hydrophobicity, and interactions within multispecies communities are key determinants of L. monocytogenes persistence on food-contact surfaces. These properties influence bacterial attachment, biofilm maturation, and resistance to mechanical removal by modulating adhesion and the stability of the extracellular polymeric substances (EPS) matrix [[18], [19], [20], [21]]. Understanding how these characteristics are affected by both native environmental microbiota and surface material is essential for evaluating biofilm behaviour under food-processing conditions.

Cross-contamination during food processing is a critical route for the transmission of foodborne pathogens from contaminated surfaces to food products. During postharvest operations such as washing, cutting, and conveying, leafy vegetables come into repeated contact with equipment and surfaces, facilitating the transfer of L. monocytogenes from environmental reservoirs to edible tissues [[22], [23], [24]]. The likelihood of pathogen transfer depends not only on the presence of L. monocytogenes on food contact surfaces, but also on its ability to detach and be transmitted during contact events. Transfer efficiency is influenced by factors such as surface material, moisture, contact conditions, and food characteristics [[25], [26], [27]]. In fresh produce systems, high surface hydration and the irregular structure of plant tissues may further favour bacterial attachment following transfer. Biofilms substantially modify cross-contamination dynamics compared to planktonic cells. Cells embedded within biofilms are retained within an extracellular polymeric matrix (EPS) that can modulate detachment behaviour and alter cell surface properties, such as hydrophobicity and adhesion forces, thereby influencing transfer efficiency [28,29].

This study aimed to determine how native environmental microbiota and surface material influence L. monocytogenes behaviour in biofilms formed on food-contact surfaces. Using a low-shear drip-flow model, mono- and multispecies biofilms were compared on the selected surfaces by evaluating biofilm development, transfer and residual retention on spinach leaves, cell surface hydrophobicity, and biofilm organization.

2. Material and methods

2.1. Bacterial culture and inoculum preparation

Native microbiota (NEM) were collected at one fresh-cut processing facility and stored at −80 °C [9]. Five L. monocytogenes strains, FS2013 (serotype 4b, milk outbreak), FS2017 (serotype 4b, beef and pork outbreak), FS2025 (serotype 1/2b, cantaloupe outbreak), FS2063 (serotype 4b, cheese outbreak), and FS2064 (serotype 1/2a, celery outbreak), were used as a cocktail for biofilm growth and for cell enumeration. FS2063 and FS2064 carried chromosomally encoded green fluorescence protein (GFP) and were used for microscopic examinations of biofilms. L. monocytogenes stock cultures were maintained at −80 °C and activated by streaking on Harlequin Listeria Chromogenic Agar (HLCA) (Neogen, Lansing, MI), then incubated at 37 °C for 48 h. For inoculum preparation, one colony per strain and 100 μL of glycerol-preserved NEM sample were inoculated in Tryptic Soy Broth (TSB; BD Biosciences xxx) and incubated overnight at 37 °C with shaking (150 rpm). Cells were harvested by centrifugation (4500 × g, 5 min), washed in 0.9% saline, and adjusted to OD600 = 0.2 for L. monocytogenes and 0.1 for NEM (∼8 log CFU/mL).

2.2. Biofilm formation under low-shear conditions

Drip-flow biofilm reactors (BioSurface Technologies, Bozeman, MT) were used to simulate low-shear flow. Stainless-steel (SS), rubber silicone (RS) and polycarbonate (PC) food contact surfaces, that are commonly used in the food processing industry, were used as abiotic solid substrata for biofilm formation. Coupons (75 mm × 25 mm) were soaked in an alkali detergent for 1 h and washed thoroughly with sterile deionized water and autoclaved at 121 °C for 15 min before each use. Individual coupons were placed in the reactor holding chambers and submerged in 15 mL of TSB inoculated with overnight culture of L. monocytogenes or 1:1 of L. monocytogenes and NEM mixture to approximately 10⁶ CFU/mL. After a 5 h incubation at room temperature (RT, 22 °C), the inoculated medium was discharged, and 1% of TSB were continuously delivered to each coupon at a rate of 0.83 mL/min using a Masterflex peristaltic pump (Cole Parmer, Vernon Hills, IL). The chambers hosting the coupons were tilted to 10° to facilitate steady flow of fresh medium across the coupon surfaces and to the draining outlet. Biofilm growth consisted of two cycles of 18 h nutrient supply and 6 h drying (total of 48 h). Coupons were removed from the reactors after the 5 h initial attachment and after completion of the 48 h biofilm formation cycles for cell enumeration or other subsequent experiments.

2.3. Transfer biofilm cells from food contact surface coupons to spinach surfaces

Baby spinach leaves were purchased from a local supermarket. Both sides of the leaves were irradiated with UV lamps for 15 min to eliminate detectable indigenous microbiota. A modified version of the transfer method described by Wang et al. [26] was used to evaluate the cross-contamination potential of L. monocytogenes. Briefly, a spinach leaf was placed on the surface of each coupon with mature (48 h) biofilms. To facilitate even contact, a sterile stainless-steel coupon (27.44 ± 0.03 g) was placed on top of the spinach, which generated a static pressure of 144 Pa, for 1 min. This transfer procedure was repeated sequentially five times using fresh spinach leaves for each coupon. For each contact, the number of cells transferred (L. monocytogenes and NEM) to the spinach leaves was determined by plate count enumeration.

2.4. Enumeration of attachment cells

For cell enumeration on coupons, after initial attachment and 48 h biofilm growth, each coupon was gently washed by immersion in 20 mL of sterile 0.9% saline buffer for 1 min at 180 rpm, then the biofilm side of each coupon was carefully swabbed, followed by 5 min of sonication and 10 s of vortexing to release swab cotton-bound cells into 1 ml of 0.9% saline. For cell enumeration on spinach leaves, individual leaves were transferred into sterile stomacher bags containing 10 mL of 0.9% saline and processed for 2 min using a stomacher. The resulting suspension was then centrifuged at 4500 × g for 5 min and the pellet was resuspended in 1 mL of 0.9% saline. Cell suspensions were 10-fold serially diluted and plated on HLCA plates for enumerating L. monocytogenes populations, and on TSA supplemented with 100 μg/mL 5-Bromo-4-Chloro-3-Indolyl β-D-Glucuronide (X-Gluc) (Chem-Impex, Wood Dale, IL) for total aerobic counts. NEM populations were inferred from the total aerobic counts by subtracting counts of characteristic β-glucuronidase expressing compact blue L. monocytogenes colonies.

2.5. Determination of hydrophobicity of biofilm cells

Cell surface hydrophobicity (CSH) of biofilm cells was assessed by microbial adhesion to hydrocarbon (MATH), based on affinity to the non-polar solvents of xylenes and n-hexadecane [18], with minor modifications. Briefly, the mono- or multispecies biofilm cell suspension from different food contact surfaces were washed twice by centrifugation (7000 rpm, 10 min, 4 °C) and resuspended in 5 mL of sterile PBS. This cell suspension (2 mL) was overlayed with 1 mL of n-hexadecane or xylene. After vigorous vortexing for 2 min, the mixtures were allowed to separate for 1 h at room temperature. The CSH of biofilm cells were determined as the percentage of cells that were excluded from the aqueous phase, based on the equation:

$$CSH\%=\left(1-\frac{A1}{A0}\right)\times 100\%$$

where A0 is the optical density (OD₆₀₀) of the initial suspension, and A1 is that of the aqueous phase after solvent extraction.

2.6. Fluorescence microscopy analysis of biofilms

Fluorescence microscopy was performed using a Cytation 5 Cell Imaging Multimode Reader (Agilent, USA). For imaging assays, GFP-expressing L. monocytogenes strains FS2063 and FS2064 were used in mono- and multispecies biofilms in combination with NEM. Biofilms were developed on coupons under the drip-flow conditions described above and imaged after completion of the 48h biofilm formation period. Prior to imaging, biofilms were stained with SYTO 61 red fluorescent nucleic acid stain (Invitrogen, xxx, USA) prepared in sterile deionized water according to the provider's instructions. Staining was performed in the dark for 20 min, followed by gentle rinsing with sterile deionized water to remove excess dye. Coupons were imaged using appropriate filter sets to detect GFP-expressing L. monocytogenes and SYTO 61-stained cells representing the total microbial population.

2.7. Statistical analysis

Data obtained from three independent experiments were analysed statistically using SigmaPlot® 13.0 (Systat Software, USA). Normality and homogeneity of variance were verified prior to analysis. One-way ANOVA was used to evaluate differences between mono- and multispecies biofilms within each surface. For comparisons among surfaces, one-way ANOVA followed by Tukey's HSD post hoc test was applied. When only two groups were compared within the same surface, pairwise t-tests with Bonferroni correction were used to account for multiple comparisons. Statistical significance was set at P < 0.05.

3. Results

3.1. Biofilm development on different food contact surfaces

L. monocytogenes biofilm formation was evaluated on SS, SR, and PC food contact surfaces under mono- and multispecies conditions at initial attachment (5 h) and mature biofilm (48 h) (Fig. 1). At 5 h (1A), monospecies L. monocytogenes biofilms showed similar populations across surfaces, with mean values of 3.85 ± 0.31, 3.10 ± 0.32, and 3.05 ± 0.16 log CFU/coupon on SS, SR, and PC, respectively, whereas multispecies conditions resulted in significantly higher counts, reaching 6.42 ± 0.12 and 5.33 ± 0.12 log CFU/coupon on SS and SR, while no significant difference on PC. After additional 48 h cycles of biofilm growth (1B), L. monocytogenes counts significantly increased for both mono- and multispecies biofilms on all tested surface, while the largest log increase were seen with mono-species biofilms on SS and SR, reaching 6.11 and 5.31 log, respectively. Nevertheless, L. monocytogenes counts in multispecies biofilms were significantly (on SS) or marginally (on SR and PC) higher than those in monospecies biofilms, reaching 8.60 ± 0.16, 6.33 ± 0.08 and 5.51 ± 0.08 log CFU/coupon on corresponding surfaces. Two-way ANOVA revealed a significant interaction between surface material and biofilm type at both the initial attachment (5 h) and mature biofilm (48 h) stages (P < 0.05), indicating that the effect of NEM on L. monocytogenes persistence depended on the surface material. NEM populations, on the other hand, was largely not affected by co-inoculation with L. monocytogenes (with exception of PC) for the initial attachment (1C). After 48 h, total NEM populations were significantly higher in co-inoculated biofilms than in NEM-only biofilms, particularly on SS and SR surfaces (1D).

Fig. 1.

Formation of Listeria monocytogenes (A, B) monospecies and multispecies biofilms and native environmental microbiota (NEM) biofilms (C, D) on food contact surfaces under low-shear drip-flow conditions. Biofilms were quantified after initial attachment (5 h; A, C) and after 48 h of biofilm development (B, D). SS, stainless steel; SR, silicone rubber; PC, polycarbonate. White bars represent monospecies biofilms, whereas grey bars represent multispecies biofilms formed in the presence of L. monocytogenes and NEM. Data are shown as mean ± SD of three independent experiments (n = 3). Asterisks indicate significant differences between mono- and multispecies biofilms for a given surface (p < 0.05).

3.2. Transfer of L. monocytogenes from biofilms to fresh produce surfaces

The transfer of L. monocytogenes from mature biofilms to spinach surfaces was evaluated over five consecutive contacts under mono- and multispecies conditions (Fig. 2). On SS (Fig. 2A), L. monocytogenes transfer from monospecies biofilms gradually declined from 3.18 ± 0.04 to 1.82 ± 0.32 log CFU/leaf. In contrast, transfer from multispecies biofilms was initially lower but increased across successive contacts, reaching 3.21 ± 0.04 log CFU/leaf by the fifth contact. NEM transfer from multispecies biofilms with L. monocytogenes was significantly higher than that of L. monocytogenes during the initial contacts, but exhibited a declining pattern, dropping to a level lower than that for L. monocytogenes by the fifth contact. For biofilms on SR, L. monocytogenes from both mono- and multispecies biofilms transferred at a high level of approximately 4.2 log CFU/leaf on the first contact, followed by rapid (monospecies) or gradual (multispecies) declining. While L. monocytogenes transfer from multispecies remained detectable across all five contacts, ranging from 4.29 ± 0.01 to 2.40 ± 0.12 log CFU/leaf, that from monospecies biofilms dropped to below the detection limit by the third contact. NEM transfer also dropped to below detection by the fourth contact. On PC, both mono- and multispecies biofilms transferred quantifiable levels of L. monocytogenes only during the first contact, with values of 3.62 ± 0.16 and 4.29 ± 0.11 log CFU/coupon, respectively, and remained detectable during the second contact, but decreased sharply below the detection limit by the third contact. NEM followed a similar pattern, with transfer limited to the first two contacts.

Fig. 2.

Sequential transfer of Listeria monocytogenes and native environmental microbiota (NEM) from mature biofilms to spinach surfaces over five consecutive contacts on stainless steel (SS) (A), silicone rubber (SR) (B), and polycarbonate (PC) (C). Yellow, orange, green, blue, and purple bars represent the cumulative transferred populations after the first, second, third, fourth, and fifth sequential contacts, respectively. The red dashed line indicates the detection limit. Data are shown as mean ± SD of three independent experiments (n = 3). Different letters indicate statistically significant differences among conditions within the same transfer number (p < 0.05).

3.3. Hydrophobicity of biofilm cells assessed by MATH assay

Cell surface hydrophobicity (CSH) of biofilm-associated cells was assessed using the microbial adhesion to hydrocarbon (MATH) assay with xylenes and n-hexadecane as non-polar solvents (Fig. 3). In the xylenes assay (3A), multi-species biofilms on SS showed the highest hydrophobicity, averaging 41.48 ± 3.71%, significantly greater than the 7.87 ± 3.56% observed for L. monocytogenes mono-species biofilms. A similar trend was observed on SR, with values of 20.26 ± 0.76% and 4.91 ± 1.10% for multi- and mono-species biofilms, respectively. On PC, hydrophobicity was also higher under multispecies conditions, 16.34 ± 7.08%, compared to monospecies biofilms, 6.71 ± 3.32%. Overall, SS supported the highest hydrophobicity under multispecies conditions, followed by SR and PC. In the n-hexadecane assay (4B), the same general trend was observed on SS, where multispecies biofilms exhibited higher hydrophobicity, averaging 34.73 ± 12.17%, compared to 17.98 ± 9.89% for mono-species biofilms, although no significant differences were detected. On SR, hydrophobicity values were also comparable between mono- and multispecies biofilms. In contrast, on PC, multispecies biofilms showed significantly higher hydrophobicity, 29.21 ± 2.06%, compared to monospecies biofilms, 23.66 ± 1.42%.

Fig. 3.

Cell surface hydrophobicity (CSH) of biofilm-associated cells determined by the microbial adhesion to hydrocarbon (MATH) assay using xylenes (A) and n-hexadecane (B). White bars represent Listeria monocytogenes monospecies biofilms, and grey bars represent multispecies biofilms composed of L. monocytogenes and native environmental microbiota (NEM). CSH is expressed as percentage of cells excluded from the aqueous phase. Data are presented as mean ± SD of three independent experiments (n = 3). Different letters indicate statistically significant differences between mono- and multispecies biofilms on the same surface, whereas asterisks denote significant differences among surfaces within the same biofilm condition (p < 0.05).

3.4. Surface-dependent biofilm organization by fluorescence microscopy

Fluorescence microscopy was used to qualitatively assess biofilm formation on SS, SR, and PC surfaces after 48 h under mono- and multispecies conditions (Fig. 4). Clear differences in surface coverage and spatial organization were observed depending on both surface material and biofilm composition. On SS, monospecies biofilms of L. monocytogenes showed limited surface coverage, with cells appearing sparsely distributed. In contrast, multispecies biofilms displayed higher biomass, characterized by the presence of small, discrete clusters distributed across the surface, in which L. monocytogenes appeared closely associated with the surrounding microbial population. On SR, biofilms exhibited a more homogeneous distribution compared to SS. Monospecies biofilms showed moderate and relatively uniform coverage, while multispecies biofilms appeared more continuous, with increased overall signal intensity and a more evenly distributed cell population. On PC, monospecies biofilms showed very limited surface coverage, with only scattered cells visible. Under multispecies conditions, biofilms were overall more uniformly distributed, but also with large well-defined clusters.

Fig. 4.

Visualization of L. monocytogenes and NEM in mono- and multispecies biofilms on stainless steel, silicone rubber and polycarbonate coupons at different stages of biofilm development. GFP-expressing L. monocytogenes strains FS2063 and FS2064 are visualized alongside total biomass stained with SYTO™ 61 (red). Overlapping signals (appearing yellow to orange) indicate co-localization or proximity of GFP-expressing L. monocytogenes cells within the total biomass of the native environmental microbiota (NEM).

4. Discussion

This study demonstrated that L. monocytogenes persistence was influenced by both surface material and the presence of NEM. Across all tested conditions, SS supported the highest L. monocytogenes levels, followed by SR, while PC consistently showed the lowest values. These observations are consistent with previous studies indicating that SS provides favourable conditions for biofilm development due to its moderate roughness and surface energy, which promote cell adhesion and EPS stability [30]; [31].

The enhanced persistence of L. monocytogenes in multispecies biofilms, particularly on SS, is associated with the establishment and maturation of NEM under the same conditions. Higher NEM densities in multispecies and mature biofilms indicate the development of a structured matrix, which may contribute to L. monocytogenes persistence through increased EPS production, altered nutrient gradients, and physical shielding [32]; [9]. While some multispecies communities can be expected to inhibit L. monocytogenes through competitive exclusion or bacteriocin production [33], the results here suggest that complex native microbiota may instead promote pathogen persistence. This observation is consistent with recent studies showing that dual-species biofilms involving L. monocytogenes and Pseudomonas spp. can alter biofilm biomass, stress tolerance, antimicrobial resistance, and pathogen persistence under food-relevant conditions [[34], [35], [36], [37]]. However, unlike these defined dual-species systems, the present study used native environmental microbiota, indicating that resident microbial communities can also modulate L. monocytogenes persistence in a surface-dependent manner. SR also supported elevated L. monocytogenes levels under multispecies conditions, indicating that surface hydrophobicity alone does not dictate persistence. Rather, the combined effects of surface properties, microbial interactions, and matrix development appear to govern colonization [38,39]. In contrast, lower NEM and L. monocytogenes populations on PC suggest reduced biofilm stability on smoother, low-energy surfaces [40,41]. Overall, these observations indicate that L. monocytogenes persistence on food-contact surfaces is driven not only by surface characteristics but also by the structure and development of the surrounding microbial community, highlighting the importance of considering native microbiota when assessing contamination risks.

The risk of L. monocytogenes cross-contamination during food processing is closely linked to the transfer potential of cells from established biofilms, which is modulated by both biofilm composition and surface material. On SS, monospecies biofilms released high levels of L. monocytogenes during the initial contacts, followed by a gradual decline, whereas multispecies biofilms showed a progressive increase in transfer across successive contacts. This observation is consistent with a stratified biofilm structure, in which NEM may dominate the outer layers while L. monocytogenes remains more protected within the matrix, as previously demonstrated by CLSM imaging [9]. Similar spatial structuring has been described in complex communities, where interactions and EPS architecture influence cell positioning and biofilm resistance [42]; [20,35,39]. Surface material also played a major role in determining transfer behaviour. Biofilms on SR surface supported sustained L. monocytogenes transfer over multiple contacts under multispecies conditions, consistent with a more homogeneous biofilm distribution and stable adhesion [43]; [44]. In contrast, those on PC surface exhibited a distinct pattern, with high L. monocytogenes release during the first contact, followed by a rapid decline to undetectable levels. This behaviour is consistent with its low surface energy and smoothness, which reduce microbial retention and facilitate detachment under mechanical stress [40,45]. Altogether, these observations highlight that while biofilm composition influences internal organization and release dynamics, surface properties largely determine the extent and persistence of cell transfer during contact events, with direct implications for cross-contamination risk. However, compared to data in Fig. 1, the cumulative transferred populations of L. monocytogenes and NEM after the five contacts constituted a minor proportion of the initial biofilms. This was true events for biofilms on PC surface, where cell transfers after the initial two contacts were reduced to below the detection limit, indicating large biofilm populations were more closely attached to the biofilm matrices and the substrata, which would serve as a persistent source of contamination. Caution is needed with this interpretation, as UV-C exposure was shown to reduce L. monocytogenes attachment on cut lettuce pieces [46]. We did not observe evidence that UV treatment used to reduce background microbiota on spinach leaves in the transfer assays induced physiological responses that would influence L. monocytogenes attachment or survival, nonetheless, this potential warrants further testing.

Differences in transfer dynamics may also be influenced by the surface properties of biofilm-forming cells. In particular, cell surface hydrophobicity (CSH) affects microbial adhesion and biofilm stability, especially in multispecies systems [19,44]. In this study, CSH was assessed using the MATH assay with xylene and hexadecane, which interact with different regions of the cell envelope [47]. Overall, CSH was higher in multispecies biofilms, particularly on SS, suggesting that NEM may enhance hydrophobic character, potentially through changes in EPS composition [48]; [49]. In contrast, CSH values remained within a similar range between mono- and multispecies biofilms on SR and PC, indicating a more limited effect of microbial interactions under these conditions. These findings suggest that both microbial composition and surface material influence the expression or exposure of hydrophobic domains [21,50]. Higher CSH values were generally observed with xylene than with hexadecane, indicating a greater contribution of surface-exposed hydrophobicity to adhesion. Notably, the elevated CSH of multispecies biofilms on SS is consistent with their higher persistence and retention, whereas the lower CSH observed on PC aligns with its reduced biofilm stability and retention [40,41]. Together, these results support the role of CSH as a contributing factor to biofilm resilience and highlight the need to consider both microbial interactions and material properties when assessing pathogen persistence.

This study underscores the importance of considering both microbial composition and surface type when evaluating L. monocytogenes persistence. Fluorescence microscopy provided additional insight into biofilm organization across surfaces. Multispecies biofilms were characterized by clustered arrangements of L. monocytogenes, with smaller, more distributed aggregates on SS and larger, localized clusters on PC, while SR showed a more homogeneous distribution. These structural patterns are consistent with previous observations on SS, where L. monocytogenes formed discrete clusters associated with NEM biomass and occupied protected niches within the biofilm [9]. Similar spatial organization has been reported in complex biofilms, where matrix structure and microbial interactions drive cell positioning [42]; [20,35]. The observed structural differences likely reflect variations in surface properties and matrix organization. The SS biofilms displayed more heterogeneous and clustered structures, suggesting enhanced structural cohesion, whereas SR and PC showed more homogeneous fluorescence coverage. These observations may explain the apparent differences between fluorescence microscopy and CFU-based analyses, as microscopy reflects total biofilm coverage and spatial organization, whereas CFU enumeration measures cultivable and transferable populations.

Although this study demonstrates that NEM altered L. monocytogenes behaviour in a surface-dependent manner, the specific microbial members and molecular mechanisms driving these effects remain to be determined. Future work integrating taxonomic profiling, biofilm architecture analysis, and functional approaches would help clarify how native communities shape pathogen persistence and detachment on different food-contact materials.

5. Conclusion

This study demonstrates that L. monocytogenes persistence, transfer dynamics, and cell surface hydrophobicity are influenced by both native microbiota and the physicochemical properties of food-contact surfaces. Multispecies biofilms promoted higher pathogen persistence and sustained transfer during repeated contact events, particularly on SS, whereas PC showed reduced biofilm stability and limited long-term persistence. These patterns were associated with differences in biofilm organization and cohesion under multispecies conditions. Importantly, the results indicate that surface type governs not only biofilm formation but also subsequent cross-contamination behaviour.

CRediT authorship contribution statement

Irene Falco: Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Writing – original draft. Yishan Yang: Writing – review & editing. Jitendra Patel: Writing – review & editing. Xiangwu Nou: Conceptualization, Supervision, Writing – review & editing.

Declaration of competing interest

The authors declare the following financial interests/personal relationships which may be considered as potential competing interests:Xiangwu Nou reports financial support was provided by US Department of Agriculture. If there are other authors, they declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Data availability

Data will be made available on request.