The study was organized in seven experimental phases: (1) definition of the dual-species biofilm model with quantification of biomass production and metabolic activity on polystyrene and silicone elastomer and comparison with the corresponding single species counterpart; (2) identification of the non-cytotoxic BSs concentrations with a significant inhibitory activity against single species biofilm formation; (3) evaluation of the anti-biofilm and antimicrobial activity of BSs at the non-cytotoxic concentrations against polymicrobial cultures, in co-incubation; (4) evaluation of the anti-biofilm and antimicrobial activity of silicone discs coated with BSs against polymicrobial cultures; (5) assessment of cells surface hydrophobicity and membrane permeability changes induced by BSs in soluble form; (6) observation of the dual-species biofilm micro-structure on BSs-coated silicone discs; and (7) preliminary assessment of BSs-coated silicone discs biocompatibility.

The rhamnolipid-producer strain Pseudomonas aeruginosa 89, a clinical isolate from a patient with cystic fibrosis, was cultured from frozen stocks onto Tryptic Soy Agar (TSA, Scharlab, Barcelona, ES) at 37°C for 18–20 h. The lipopeptide-producer strain Bacillus subtilis AC7, from the inside of stems of Robinia pseudoacacia, was cultured from frozen stocks onto Luria Bertani agar (LBA, Sigma–Aldrich, St. Louis, MO, United States) at 28°C for 18–20 h. The sophorolipid-producer strain Candida bombicola ATCC 22214, obtained from the American Type Culture Collection (ATCC, Manassas, VA, United States), was cultured from frozen stocks onto Sabouraud dextrose agar (SDA, Scharlab, Barcelona, ES) at 25°C for 18–20 h. All the biofilm-producer strains used in this study were obtained from the ATCC (Manassas, VA, United States). C. albicans ATCC 10231, S. aureus ATCC 25923, S. aureus ATCC 6538, and S. epidermidis ATCC 35984 were cultured from frozen stocks onto SDA and TSA plates, respectively, and incubated overnight at 37°C.

Lipopeptide AC7BS and rhamnolipid R89BS were produced and extracted as described by Ceresa et al. (2018, 2019b). Briefly, a loop of B. subtilis AC7 from a LBA overnight culture was grown in 20 ml of LB broth at 28°C for 4 h at 140 r/min. Afterward, 2 ml of this culture was used to inoculate 500 ml of LB broth and was incubated at 28°C for 24 h at 140 r/min. A loop of P. aeruginosa 89 from a TSA overnight culture was grown in 40 ml of Nutrient Broth II (Sifin Diagnostics GmbH, Berlin, DE) for 4 h at 37°C at 140 r/min. Afterward, 8 ml of this culture was added to 400 ml of Siegmund–Wagner medium and incubated at 37°C for 5 days at 120 r/min. Cell-free supernatants were acidified to pH 2.2 with 6 M HCl (AC7BS) or 6 M H₂SO₄ (R89BS) and stored overnight at 4°C. Lipopeptide AC7BS and rhamnolipid R89BS were extracted three times with 167 ml ethyl acetate:methanol (4:1) or 134 ml ethyl acetate (Sigma–Aldrich, St. Louis, MO, United States), respectively. Organic phases were anhydrified, filtrated, and vacuum-dried. BSs were recovered by dissolution in acetone (Sigma–Aldrich, St. Louis, MO, United States) and collected in glass tubes. Acetone was, then, evaporated and BSs were weighted.

Sophorolipid SL18 was obtained from a fed batch cultivation of C. bombicola ATCC 22214, according to Ceresa et al. (2020). Briefly, the cells (10% v/v) were grown in 2 l of Glucose Yeast Urea (GYU) medium [100 g/l glucose (Sigma–Aldrich, St. Louis, MO, United States), 10 g/l yeast extract (Sigma–Aldrich, St. Louis, MO, United States), and 1 g/l urea (Sigma–Aldrich, St. Louis, MO, United States)]. Oleic acid (99%, Sigma–Aldrich, St. Louis, MO, United States) was supplemented as a feeding source at a concentration of 20% to generate lactonic congeners. Fermentation was performed for 8 days at 200 r/min and 30°C. Sophorolipid SL18 were, then, extracted twice with ethyl acetate (1:1 extract ratio) (Sigma–Aldrich, St. Louis, MO, United States) and partially purified by three washings with hexane (Sigma–Aldrich, St. Louis, MO, United States) to remove residual fatty acids.

At the end of the extraction process, the presence, the purity, and the composition of the three BSs were confirmed by ESI/MS analysis as previously described (Ceresa et al., 2016, 2019b, 2020). All the following biological tests and microscopy analyses were performed using the same batch of production for each BS. BSs were dissolved in Phosphate Buffer Solution pH 7.4 (PBS) at the different concentrations of use. The solutions were filtered through a 0.2 μm filter and stored at room temperature.

Silicone-elastomeric discs (SEDs—0.8 cm in diameter and 1.5 mm in thickness) were cut from medical-grade silicone sheets (TECNOEXTR s.r.l, Palazzolo sull’Oglio, IT) and prepared as described in Ceresa et al. (2015). Briefly, discs were cleaned with a 1.4% (v/v) RBS^TM 50 solution (Sigma–Aldrich, St. Louis, MO, United States), sonicated for 5 min at 60 kHz, and rinsed twice in Milli-Q water. Silicone was, then, dipped in MeOH (99%, Sigma–Aldrich, St. Louis, MO, United States), sonicated, and rinsed as previously described. Afterward, SEDs were autoclaved, dried, and moved aseptically into 48-well plates.

Mono- and dual-species biofilm formation: fungal and bacterial cells were suspended in Roswell Park Memorial Institute (RPMI) 1640 (Sigma–Aldrich, St. Louis, MO, United States) buffered with MOPS (Sigma–Aldrich, St. Louis, MO, United States) and supplemented with 2% Glucose (Biolife, Monza, IT), pH 7.0 (RPMI +2%G). Cell density was adjusted up to 10⁶ and 10⁷ colony forming unit (CFU)/ml for C. albicans and Staphylococcus spp. respectively. Polystyrene was used as a substrate for the growth of biofilms in co-incubation assays and silicone was used for the growth of biofilms in the coating assays. Surfaces were inoculated with 0.5 ml of the suspension and incubated at 37°C in static conditions up to 72 h; growth medium was removed and replaced with fresh RPMI +2%G every 24 h. Blank polystyrene and silicone control surfaces (without biofilm) were also included in the experimental setting. The ability of microbial strains to form polymicrobial biofilms, compared to the mono-species ones, was evaluated by the determination of biofilm biomass and metabolic activity of sessile cells as described below. All experiments were carried out in quadruplicate and repeated two times.

Co-incubation conditions (BSs in soluble form in polystyrene plates): in order to determine the minimum non-cytotoxic concentration of BSs (Ceresa et al., 2018, 2019b) able to inhibit single-species biofilm formation on polystyrene by at least 80%, increasing concentrations of lipopeptide AC7BS (0.125–0.5 mg/ml) and rhamnolipid R89BS (0.0125–0.05 mg/ml) were tested. The wells were filled with 50 μl of 10× BSs solutions (treated samples) or with an equal volume of PBS (control samples) and 0.5 ml of single-species suspensions of C. albicans (10⁶ CFU/ml) and Staphylococcus spp. (10⁷ CFU/ml) in RPMI +2%G. The 48-well plates were incubated for 24 h at 37°C. The effect was evaluated in terms of biofilm biomass reduction as described below. All experiments were carried out in quadruplicate and repeated twice.

Subsequently, the selected concentrations of BSs were tested against dual-species biofilm formation. The bottom of the wells was covered with 50 μl of the selected 10× BSs solutions (treated samples) or with an equal volume of PBS (control samples). Then, 0.5 ml of the dual-species suspensions of C. albicans (10⁶ CFU/ml) and Staphylococcus spp. (10⁷ CFU/ml) in RPMI +2%G were added to each well. The plates were incubated up to 72 h at 37°C. Growth medium was removed and replaced with fresh RPMI +2%G supplemented with 10× BSs solutions (treated samples) or with an equal volume of PBS (control samples) every 24 h. Blank surfaces (without biofilm) were also included in the experimental setting. The ability of microbial surfactants to inhibit dual-species biofilm formation in co-incubation conditions was evaluated by the determination of biofilm biomass and metabolic activity of sessile and planktonic cells as described below. Experiments were performed in quadruplicate and repeated twice.

Coating conditions: the treatment of silicone surfaces was carried out in 48-well plates by immersing SEDs in BS solutions (rhamnolipid R89BS: 2 mg/ml; lipopeptide AC7BS: 2 mg/ml; sophorolipid SL18: 8 mg/ml) at 37°C for 24 h at 180 r/min. These solutions were chosen as previously optimized in Ceresa et al. (2016; 2019b; 2020). Afterward, discs were moved into new plates and dried before use. Five hundred microliters of the dual-species suspensions (C. albicans at the concentration of 10⁶ CFU/ml and Staphylococcus spp. at the concentration of 10⁷ CFU/ml) in RPMI +2%G were added to each well. SEDs were incubated up to 72 h at 37°C. Every 24 h, discs were moved into fresh media. Blank surfaces (without biofilm) were also included. The anti-biofilm activity of BSs was evaluated at 24, 48, and 72 h by the determination of biofilm biomass, metabolic activity of sessile and planktonic cells, and viable cell counting as described below. Experiments were performed in quadruplicate and repeated twice.

The number of microbial cells forming the multi-species biofilms was determined by the spread plate method as described in Ceresa et al. (2015). Briefly, after two washings, biofilms were detached from silicone surfaces and broke up by four cycles of sonication (30 s) and stirring (30 s). The obtained suspensions were serially diluted in PBS and seeded both on Mannitol Salt Agar (MSA, Scharlab, Barcelona, ES) plates, selective for staphylococcal species, and on Sabouraud Chloramphenicol Agar (SCA, Scharlab, Barcelona, ES) plates, selective for fungal species. After 24 h at 37°C, the colonies were counted and the number of C. albicans, S. aureus, or S. epidermidis cells within the polymicrobial biofilm was quantified.

Silicone-elastomeric discs surface coating with the BSs was carried out as described by Papa et al. (2015), with minor changes. Briefly, a volume of 20 μl of BS solutions (rhamnolipid R89BS: 2 mg/ml; lipopeptide AC7BS: 2 mg/ml; sophorolipid SL18: 8 mg/ml) or 20 μl of PBS as control, were deposited on the silicone surfaces. SEDs were then placed under laminar flow to allow complete drying and, subsequently, moved into 48-well plates. The discs were filled with 0.5 ml of the dual-species suspensions (C. albicans at the concentration of 10⁶ CFU/ml and Staphylococcus spp. at the concentration of 10⁷ CFU/ml in RPMI +2%G) and incubated at 37°C for 4 h. The quantification of cells attached on SEDs was carried out using crystal violet staining as reported in Section “Biofilm Biomass.” All experiments were carried out in quadruplicate and repeated twice.

The micromorphology and architecture of multi-species biofilm on SEDs was visualized using the scanning electron microscope (SEM) Quanta 200F FEG (Fei, Eindhoven, Netherlands) in high-vacuum mode. Samples were prepared for SEM imaging according to Ceresa et al. (2019b), with minor changes. Briefly, after two washings, biofilms were fixed in 2.5% glutaraldehyde, washed twice in distilled water, dehydrated in a graded ethanol series, and coated with a 10 nm layer of gold with a sputter coater (Emitech K500X, Quorum Technologies, Laughton, United Kingdom). A set of representative images at a magnification of 500×, 1000×, 2000×, and 4000× were obtained from untreated (controls) and pre-coated SEDs with rhamnolipid R89BS or sophorolipid SL18 and incubated with either C. albicans–S. epidermidis or C. albicans–S. aureus at 24, 48, and 72 h. Secondary electron signal was collected to investigate structural details of microbial cells and extracellular matrix on the biofilm. The primary beam energy was set to 5 keV to minimize damage to the organic structures. Possible artifacts due to the sample preparation process were considered according to indications provided by Hrubanova et al. (2018) and previous experience performed in imaging microbial biofilm formed in vitro on medical devices (Tessarolo et al., 2007; Signoretto et al., 2013).

The in vitro biocompatibility of BSs-coated discs was evaluated in 48-well plates by the MTT assay (Ceresa et al., 2019a). Spontaneously immortalized human skin keratinocyte—HaCaT cells (10⁴ cells/well) were seeded in Dulbecco’s Modified Eagle’s Medium (DMEM) high glucose (EuroClone, Milan, IT) supplemented with 4% FBS (EuroClone, Milan, IT), L-glutamine 200 nM (EuroClone, Italy) and 1% Pen/Strep (EuroClone, Milan, IT), and incubated at 37°C in 5% CO₂. After 24 h, growth medium was removed and replaced with the eluates obtained from BSs-coated SEDs after static release at 37°C for 24 h. Negative control consisted in cells treated with 0.5% Triton X, whereas positive control was represented by cells w/o any treatment. Fifty microliters of the MTT solution (5 mg/ml) was added into each well. Plates were then incubated for 24 and 72 h at 37°C. Formazan crystals were dissolved with 200 μl of 0.05 M HCl/isopropanol (50:1) and A570 was measured at the two time points. The percentage of cell viability was estimated using the following formula:

where

A_sample: absorbance of BSs or CTRL—samples

A_CTRL+: absorbance of positive controls.

Experiments were performed in triplicate and repeated twice.

All analyses and graphics were performed using the statistical program R, 3.6.2 (R Core Team, 2019). One-way ANOVA was applied to compare mono- and dual-species biofilms. Two-way ANOVA followed by Tukey post hoc test was used to investigate the anti-biofilm activity of BSs on dual-species biofilms and the metabolic activity of planktonic cells. To estimate log₁₀ CFU/disk from colony counts, the R package dupiR was used (Comoglio et al., 2013). Differences in the percentage composition of dual-species biofilms were investigated by two-sample t-test for equality of proportions with continuity correction. Two-sample t-test was performed to evaluate the significance of data in hydrophobicity and membrane permeability assays. One-way ANOVA followed by Tukey post hoc test was performed to evaluate the significance of data in the biocompatibility assay in comparison to positive and negative controls. Differences were considered statistically significant at p < 0.05.

Mono- and dual-species biofilms of C. albicans–Staphylococcus spp. were grown on polystyrene (Figure 1) and on silicone surfaces (Figure 2) up to 72 h. Biofilm biomass and biofilm metabolic activity were quantified at 24, 48, and 72 h. C. albicans–S. aureus biomass and metabolic activity were higher than those of the two individual species both on polystyrene and on silicone at all incubation times (p < 0.001). Concerning C. albicans–S. epidermidis, biomass and metabolic activity were always higher than those observed for the individual species, on both surfaces, up to 48 h (p < 0.001). On the contrary, at 72 h, biomass and metabolic activity of dual-species biofilms were lower than those observed for S. epidermidis biofilms, with the exception of metabolic activity on polystyrene (p < 0.001).

AC7BS and R89BS were selected to perform the co-incubation assays as they were previously reported as not cytotoxic versus MRC5 cells monolayers at the concentrations active against biofilm (Ceresa et al., 2018, 2019b). On the contrary, SL-18 was not included in this assay since it was previously detected as cytotoxic when used in soluble form at concentrations active against C. albicans and S. aureus single species biofilms (Ceresa et al., 2020). Increasing concentrations of lipopeptide AC7BS (0.125–0.5 mg/ml) and rhamnolipid R89BS (0.0125–0.05 mg/ml) were tested and the minimum concentration of BSs that counteracted single-species biofilms formation on polystyrene by at least 80% (inhibition level threshold) was identified (Figure 3). In general, the formation of C. albicans and S. aureus biofilms was reduced in a concentration-dependent manner by the two BSs while the formation of S. epidermidis biofilm was effectively inhibited (≥80%) only by rhamnolipid R89BS (Figure 3A). In particular, the three mono-species biofilms were inhibited by about 87% by rhamnolipid R89BS at a concentration of 0.05 mg/ml. Concerning lipopeptide AC7BS (Figure 3B), the threshold inhibition level was reached at a concentration of 0.5 mg/ml only for C. albicans and S. aureus biofilms (82%) but not for S. epidermidis, which had a maximum inhibition of only 27%. For this reason, this BS was excluded from the subsequent anti-biofilm assays against C. albicans–S. epidermidis dual-species biofilms.

The boxplot in Figure 4 shows the effect of rhamnolipid R89BS and lipopeptide AC7BS against C. albicans–S. aureus and of R89BS against C. albicans–S. epidermidis biofilm formation up to 72 h. In general, as confirmed by two-way ANOVA and Tukey post hoc test, for each co-culture, biofilm formation on polystyrene was significantly dependent on BSs treatment (p < 0.001) and incubation time (p < 0.001). Regardless of the two strain combinations involved in the dual-species biofilm development, biofilm biomass (Figures 4A,D) and metabolic activity (Figures 4B,E) were equally inhibited by the BSs. The anti-biofilm activity of rhamnolipid R89BS was stable up to 72 h, while, for lipopeptide AC7BS, a slight reduction was detected between 24 and 72 h. In particular, as observed in Table 1, rhamnolipid R89BS proved to be the most effective BS for the inhibition of C. albicans–S. aureus biofilm growth and an excellent agent for the prevention of C. albicans–S. epidermidis biofilm formation, with mean percentages of reduction of 94% (C. albicans–S. aureus) and 95% (C. albicans–S. epidermidis), after 72 h co-incubation. In addition, to define whether part of the observed effect was the result of an antimicrobial activity of the BSs, the metabolic activity of planktonic cells in the supernatants was assessed (Figures 4C,F). The absorbance values of the cell supernatants co-incubated with BSs were significantly higher in comparison to the controls (p < 0.001) suggesting that, in the treated wells, cells existed in a planktonic state rather than by forming a biofilm. However, in the case of C. albicans–S. aureus co-cultures, the lower absorbance values observed in the rhamnolipid R89BS-treated wells compared to those of the lipopeptide AC7BS-treated wells (p < 0.001), suggested an antimicrobial activity of the rhamnolipid (Figure 4C).

The results of BSs pre-coating on SEDs further confirmed the inhibitory activity of these natural microbial molecules against the formation of C. albicans–S. aureus (rhamnolipid R89BS, lipopeptide AC7BS, sophorolipid SL18) and C. albicans–S. epidermidis (rhamnolipid R89BS, sophorolipid SL18) mixed biofilms (Figure 5). As previously observed in co-incubation conditions, for each co-culture, the development of biofilms on silicone surfaces was significantly dependent on BSs treatment (p < 0.001) and incubation time (p < 0.001). Biofilm biomass (Figures 5A,D) and metabolic activity (Figures 5B,E) were equally inhibited on all BSs-coated discs (SEDs). Concerning the dual-species biofilms of C. albicans–S. aureus, the anti-biofilm activity of rhamnolipid R89BS- and sophorolipid SL18-coated SEDs was stable up to 72 h, while a slight reduction of the efficacy of lipopeptide AC7BS-coated SEDs was observed during this time. The anti-biofilm activity of rhamnolipid R89BS coating was stable also for C. albicans–S. epidermidis mixed biofilms, while that of sophorolipid SL18-coated SEDs slightly decreased over time. In general, starting from 48 h of incubation, the surface treatment of silicone with rhamnolipid R89BS proved to be the most effective in counteracting the growth of both polymicrobial biofilms. In particular, after 72 h, average inhibitions of 93 and 90% against C. albicans–S. aureus and C. albicans–S. epidermidis biofilms were found, respectively. Table 2 shows the percentages of biomass and metabolic activity inhibition at the different time-points. To exclude that the observed activity was due to an antimicrobial action of the BSs, the metabolic activity of the planktonic cells in the wells was evaluated (Figures 5C,F). As observed in co-incubation conditions, the absorbance values of the supernatants of the BSs-coated silicone discs were significantly higher than those obtained for the corresponding controls (p < 0.001). Conversely, no significant variations were found between the absorbance values of planktonic cells recorded for the different BSs treatments (p > 0.05).

In addition, to evaluate whether the tested BSs exhibited their anti-biofilm action in equal proportions on the single species forming the polymicrobial biofilms on the silicone surfaces, the number of cells of C. albicans, S. aureus, and S. epidermidis was determined on selective media for both control and coated discs (Supplementary Tables 1, 2) and the percentage compositions were calculated (Tables 3, 4). In general, at all incubation times, biofilms on the control discs mainly consisted of bacterial cells (98.6–99.8%) and only for a small percentage of the yeast cells (0.2–1.4%). In particular, the yeast cells were present in a major proportion in C. albicans–S. aureus biofilms (Table 3) than in C. albicans–S. epidermidis (Table 4). Interestingly, at 24 h, the presence of BSs on the silicone surface was associated with a significant increase in the yeast species percentage compared to that observed on the controls (p < 0.001). However, this value decreased over time and returned to levels similar to those observed for the controls.

Finally, to assess the activity of BSs coating on SEDs on the early phases of biofilm formation (adhesion phase), the amount of C. albicans–S. aureus and C. albicans–S. epidermidis adherent cells was evaluated after 4 h incubation by means of the CV method (Figure 6). Similarly, to biofilm formation, microbial cells adhesion to silicone surfaces was also significantly dependent on BSs treatment (p < 0.001). Concerning C. albicans–S. aureus, the highest anti-adhesive activity was observed for rhamnolipid R89BS (71%), followed by sophorolipid SL18 (64%) and lipopeptide AC7BS (51%). As to C. albicans–S. epidermidis, rhamnolipid R89BS coating proved to be the most effective with an inhibition of adhesion of 62% while for sophorolipid SL18 showed a 54% inhibition.

In order to evaluate the possible effect of BSs on the tested opportunistic pathogens in co-incubation conditions, cell surface hydrophobicity (CSH) and membrane permeability were assessed. As shown in Figure 7A, after 1 h incubation with R89BS and AC7BS, CSH and membrane permeability of C. albicans, S. aureus, and S. epidermidis were different from the untreated controls. BSs treatment induced a significant modification of CSH for all the tested strains (p < 0.05). In particular, it resulted in a decrease of CSH both for bacterial and fungal cells. Bacterial CSH decreased from 99% (mean percentage for untreated control samples) to values ranging from 38 to 71% for BSs treated samples. Fungal CSH in comparison was reduced from 44% (mean percentage for control samples) to values ranging from 31 to 37% for BSs treated samples. The changes of CSH induced by BSs were related to the microbial strain: in particular, S. aureus was the most susceptible to cell hydrophobicity changes, followed by S. epidermidis and C. albicans. The treatment with BSs also altered cell membrane permeability for all the tested strains (p < 0.05) (Figure 7B). In particular, BSs induced an increase in crystal violet uptake by bacterial cells from 46% (mean percentage for untreated control samples) to values ranging from 54 to 74% and a slight increase in the case of fungal cells from 53% (mean percentage for untreated control samples) to values ranging from 56 to 59%. Again, the changes of membrane permeability induced by BSs were related to the microbial strain: in particular, S. aureus and S. epidermidis were more susceptible to BSs than C. albicans.

Scanning electron microscopic images presenting the features of the biofilm formed on untreated (controls) and pre-coated SEDs with rhamnolipid R89BS and sophorolipid SL18 at the longest incubation time are presented in Figure 8. The SEM investigation showed the presence of both fungal hyphae and coccoid bacteria adhering to the surface of all samples. In general, there was a more pronounced spatial correlation between S. aureus and C. albicans (Figures 8A,C,E) than between S. epidermidis and C. albicans (Figures 8B,D,F), irrespective of the surface treatment. More specifically, S. aureus was prevalently found in close contact with fungal hyphae, almost completely covering the Candida mycelium at 72 h in control SEDs (Figure 8A), while S. epidermidis never realized a full coating of the C. albicans structures (Figure 8B) and was more prone to adhere directly to the SEDs surface (Figure 8D).

In agreement with the results obtained from quantitative tests of biofilm biomass performed on corresponding samples, untreated controls showed a well-structured dual-species biofilm at each time-points, having a more three-dimensional arrangement and a thicker appearance at longer incubation times (Figures 8A,B). A similar increasing trend in the number of cells at the surface was also observed for rhamnolipid R89BS and sophorolipid SL18 pre-coated SEDs. However, the number of microbial cells at the treated surface was drastically reduced in respect of controls and the large majority of the sample surface was free of cells or presented small clusters with few three-dimensional microbial aggregates at 24 and 48 h (Supplementary Figures S1, S2). This was also the case for S. aureus and C. albicans dual-species biofilm at 72 h (Figures 8C,E), but not for S. epidermidis and C. albicans dual-species biofilm where only a minor portion of the treated surface was free of cells (Figure 8D) or mainly fully covered (Figure 8F). At 72 h, some differences in C. albicans–S. epidermidis biofilms were also noted between rhamnolipid R89BS and sophorolipid SL18 pre-coated discs, with the latter presenting areas showing a mature biofilm (Figure 8F), although less structured and thick than the controls (Figure 8B). Rhamnolipid R89BS pre-coated SEDs showed only minor portions of the disc surface still free of microorganisms (Figure 8D).

No cytotoxic effect was detected on spontaneously immortalized human skin keratinocyte when exposed for 24 h to the BSs-coated SEDs eluates obtained from static release conditions. In fact, at this time-point, the viability of HaCat cells was comparable to positive controls (p > 0.05), independently to the type of BS involved in the coating procedure (Supplementary Figure S3A). After 72 h of cells exposure to the eluates, cell viability slightly decreased to values ranging from 92% for SL18 to 78% for R89BS (Supplementary Figure S3B) but was always above the limit (70%) according to the ISO 10993-5 standard (Xian, 2009; Rodríguez-López et al., 2020).