Chemicals and reagents including rhamnolipids mixture R90 (AGAE Technologies) and 4-(Dimethylamino)pyridine (DMAP) were purchased from Sigma–Aldrich (Munich, Germany). Reagents used for coupling reactions including N-(3-Dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride (EDCI) were purchased from ACROS Organics (Morris Plains, NJ, United States). Unless otherwise stated, all media components were purchased either from Oxoid (Cambridge, United Kingdom) or Becton Dickinson (Sparks, MD, United States).

Gentamycin sulfate and chloramphenicol were purchased from Sigma–Aldrich (Munich, Germany), tetracycline hydrochloride from USB Co. (Cleveland, OH, United States) and cycloheximide from SERVA (Heidelberg, Germany). The stock solutions of antibiotics were prepared as follows: gentamycin 30 g L^-1 in water, tetracycline 15 g L^-1 in ethanol, chloramphenicol 50 g L^-1 in ethanol, and cycloheximide 70 g L^-1 in water and were kept at -20°C.

Bacteria were isolated from the rhizosphere of medicinal plant Glechoma hederacea L. (ground ivy), using previously described procedure (Djokic et al., 2011). The medium for isolation of bacteria was starch casein agar containing soluble starch (10 g L^-1), casein (1 g L^-1), KH₂PO₄ (0.5 g L^-1), MgSO₄ (0.5 g L^-1), NaCl (3 g L^-1), and bacteriological agar (15 g L^-1), supplemented with antifungal cycloheximide (50 μg L^-1). Plates were incubated at 30°C for 7 days.

Isolates were grown in JS broth containing soy flour (30 g L^-1), soluble starch (20 g L^-1), glucose (20 g L^-1), mannitol (15 g L^-1), and CaCO₃ (10 g L^-1) for 7 days, at 30°C on a rotary shaker (180 rpm) and their cultures were extracted with equal volume of ethyl acetate and screened for the inhibition of biofilm formation of P. aeruginosa PAO1.

Genomic DNA isolation, PCR amplification of 16S rRNA gene sequencing and sequence alignment were performed as described previously (Stankovic et al., 2012).

Spore suspensions of BV152.1 were prepared in glycerol [20%, v/v, (Kieser et al., 2000)] (20 μl) and firstly inoculated into vegetative medium (tryptone soy broth, 8 g L^-1, yeast extract, 4 g L^-1, maltose, 15 g L^-1, and CaCO₃, 2 g L^-1) and incubated at 30°C for 48 h, and this pre-culture was then used for the inoculation of JS broth (1%, v/v). Cultures were grown in Erlenmeyer flasks (1:5 culture to volume ratio) containing coiled stainless steel spring for better aeration and incubated at 30°C on a rotary shaker (180 rpm) for 7 days.

Culture (1 L) was extracted using the equal volume of ethyl acetate by vigorous shaking for 30 min. The organic phase was then separated, dried under vacuum and further purified by flash chromatography. Flash chromatography employed silica gel 60 (230–400 mesh) and the following solvent system: n-hexane and ethyl acetate (3:1 ratio, 100 ml), n-hexane and ethyl acetate (1:1 ratio, 100 ml), ethyl acetate and methanol (9:1 ratio, 100 ml), followed by pure methanol (100 ml). Collected fractions were analyzed by thin layer chromatography, which was carried out using alumina plates with 0.25 mm silica layer (Kieselgel 60 F₂₅₄, Merck, Darmstadt, Germany) and the appropriate fractions were combined, dried under vacuum, and weighted. Anti-biofilm activity was determined for each fraction.

The fraction that showed anti-biofilm forming activity was purified further using the following solvents: pure ethyl acetate (50 mL); ethyl acetate, dichloromethane, and methanol (8:1:1 ratio, 50 mL) and ethyl acetate, dichloromethane, and methanol (4:2:1 ratio, 50 mL). The most active fraction (F3) was further analyzed by a combination of NMR and mass spectrometry. The NMR spectra were recorded on a Bruker Ascend 400 (400 MHz) spectrometer. Chemical shifts are given in parts per million (δ) downfield from tetramethylsilane as the internal standard and deuterochloroform as a solvent.

Mass spectral data were recorded using 6210 Time-of-Flight LC–MS system (Agilent Technologies, Santa Clara, CA, United States) connected to an Agilent 1200 Series HPLC instrument (Agilent Technologies, Waldbronn, Germany), with a degasser, a binary pump, an autosampler, a column compartment equipped with a Zorbax Eclipse XDB-C18 RRHT column (1.8 μm, 4.6 mm × 50 mm) and a diode-array detector, via ESI interface. The mobile phase consisted of water containing 0.2% formic acid (v/v; A) and acetonitrile (B). A gradient program was used as follows: 0–0.24 min 5% B, 0.24–10 min, 5–95% B, 10–15 min, 95% B, 15–15.5 min, 95–5% B, 15.5–20 min, 5% B. The flow rate of mobile phase was 0.5 mL/min, the column temperature was 40°C and the injection volume was 10 μL. Spectral data from all the peaks were accumulated in the range of 190–900 nm. Full scan mass spectra were measured between 100 and 1500 m/z in positive ion mode.

To a solution of di-rhamnolipids (0.1 mmol, 1 eq.) in CH₂Cl₂/DMF (2.7:0.3 mL) were added EDCI (0.12 mmol, 1.2 eq.), DMAP (0.05 mmol, 0.5 eq.) and amine (benzyl amine, piperidine and morpholine; 0.3 mmol, 3 eq.). The mixture was stirred for 16 h at room temperature. The mixture was diluted with water (10 mL) and extracted with CH₂Cl₂ (3 × 15 mL). The combined organic extract was dried with MgSO₄, filtered and concentrated in vacuo. The residue was purified by flash column chromatography (SiO₂, ethyl acetate:methanol:dichloromethane in 4:2:1 ratio) that afforded products as yellow thick oils.

To obtain silylated derivative to a solution of rhamnolipids (0.1 mmol, 1 eq.) in DMF (3 mL) TBDMS-Cl (1.6 mmol, 16 eq.) AgNO₃ (1.6 mmol, 16 eq.) and pyridine (3.2 mmol, 32 eq.) were added. The mixture was stirred overnight at ambient temperature before it was diluted with water (10 mL) and extracted with ethyl acetate (3 × 20 mL). The combined organic extract was dried with MgSO₄, filtered and concentrated in vacuum to yield the product which was purified via silica gel flash chromatography using petroleum ether/EtOAc (7:1) as eluent to provide the title compound as a colorless oil.

Physico-chemical properties of all compounds were predicted using the Marvin Sketch 17.2.13.0 program (ChemAxon¹). Consensus methods were used for calculating logP and hydrophilic-lipophilic balance (HLB) values.

Test organisms for the antibacterial assays were obtained from the National Collection of Type Cultures (NCTC) and the American Type Culture Collection (ATCC). P. aeruginosa PAO1 NCTC 10332, S. aureus ATCC 25923 and S. aureus ATCC 43300 (MRSA) and S. marcescens ATCC 27117 were used in this study. One clinical isolate of P. aeruginosa DM50 with high ability to form biofilms and resistance to metronidazole, clindamycin and amoxicillin was also included (Glisic et al., 2016). Bacterial strains were grown in Luria Bertani (LB) broth at 37°C on a rotary shaker at 180 rpm.

All rhamnolipids utilized for the bioactivity assessments in this study along with sources and descriptions are listed in Supplementary Table S1.

The minimum inhibitory concentration (MIC) of rhamnolipids and their derivatives were determined according to standard broth microdilution assays recommended by the National Committee for Clinical Laboratory Standards (M07-A8).

Stock solutions of rhamnolipids and derivatives were prepared in DMSO (50 g L^-1, w/v). The highest concentration used was 500 mg L^-1. The inoculums were 10⁵ colony forming units (cfu) mL^-1. The MIC value corresponds to the lowest concentration that inhibited the growth after 24 h at 37°C for P. aeruginosa and S. aureus or at 30°C for S. marcescens.

Biofilm quantification assays were performed in 96-well microtiter plates using a crystal violet (CV) method to stain adherent cells (Merritt et al., 2005). Overnight cultures of bacteria were diluted to 5 × 10⁷ cfu mL^-1 in LB and 100 μL was added to the wells in the presence of test compounds or DMSO (0.1%, v/v). Biofilms formed for 24 h at 37°C for P. aeruginosa and S. aureus, or 30°C for S. marcescens were washed and adherent cells stained with CV 0.1% (v/v). BFIC₅₀ (concentration of compound that inhibited biofilm formation by 50%) was determined for each compound.

The effect of rhamnolipids on P. aeruginosa biofilm formation was examined additionally by introducing 3 h adhesion phase after inoculation with 5 × 10⁷ cfu mL^-1 cells. Following adhesion, the supernatant was removed and, after two washing steps with phosphate buffered saline (PBS), test compounds or DMSO were applied and the biofilms were quantified after 24 h using CV. Each biofilm formation assay was performed in six wells and repeated at least three times.

To study the effect of di-rhamnolipids on P. aeruginosa PAO1 adherence to silicone surfaces and overnight bacterial culture was diluted to 5 × 10⁷ cfu mL^-1 in LB and 2 ml was added per well in six well microtiter plate. The silicone catheter pieces (Romed, Wilnis, Holland) of 1 cm were placed in each well containing diluted bacteria and incubated in the presence of di-rhamnolipids from Lysinibacillus sp. BV152.1 (F3) or DMSO. After 24 h, the culture medium was removed and the catheters were washed three times in PBS in order to remove the non-adherent strains. Biofilms were then fixed with cold methanol and the samples dried before the examination.

Catheters were glued to double-sided conductive carbon tab stuck on standard vacuum-clean stub, and were coated with gold (thickness of 15–20 nm) by the sputtering process (Leica EM SCD005 sputtering machine, Leica Microsystems, Mannheim, Germany). Sputtering was performed in the vacuum chamber under pressure <0.05 mbar using sputter current of 40 mA, the working distance of 50 mm and sputter time of 100 s. Such prepared samples were examined by JEOL JSM-6610LV microscope (JEOL United States, Inc., Peabody, MA, United States). An acceleration voltage of 20 kV was used.

To study the effect of di-rhamnolipids on P. aeruginosa PAO1 adherence to glass or plastic surfaces biofilms were developed on cover slips and examined under the microscope. An overnight culture of P. aeruginosa PAO1 was diluted to 5 × 10⁷ cfu mL^-1 in LB and 2 ml was added per well of six well microtiter plate containing glass or plastic cover slips. After 24 h, non-adherent cells were removed and biofilms were washed with 0.9% NaCl and stained with 2.5 μM SYTO9 green fluorescent dye and 2.5 μM propidium iodide (PI) red fluorescent dye of Live/Dead staining kit (LIVE/DEAD^® BacLight^TM Bacterial Viability Kit, Thermo Fisher Scientific, Waltham, MA, United States). Cells were observed under a fluorescence microscope (Olympus BX51, Applied Imaging Corp., San Jose, CA, United States) under 100,000 × magnification (glass) or 40,000 × magnification (plastic).

Anti-adhesion assay was performed as previously described (Shetye et al., 2014) with some modifications. An overnight culture of P. aeruginosa PAO1 containing pBBR2-GFP (da Silva et al., 2014) was subcultured (initial OD600 = 0.02) in M9+/LB (95:5) containing gentamycin (80 mg L^-1), tetracycline (20 mg L^-1), and chloramphenicol (50 mg L^-1) at 37°C in a rotary shaker (200 rpm). After reaching OD600 = 0.1, aliquots (200 μL) were transferred to the wells of black polystyrene microtiter plate with rhamnolipids, derivatives or DMSO (control). After incubation at 37°C for 2 h, bacterial cultures were discarded, and the fluorescence of adherent cells was measured after addition of fresh M9+/LB (95:5) medium using Tecan Infinite 200 Microplate reader (Tecan Group Ltd., Switzerland, lex = 500 nm, lem = 540 nm). Background signal (M9+/LB (95:5) was subtracted from all the samples. Assays were repeated three times; inhibition values are the averages of six replicate wells from one experiment.

An overnight culture of P. aeruginosa was diluted to 5 × 10⁷ cfu mL^-1 in LB and 100 μL was added to the wells in 96-well microtiter plates and incubated for 24 h at 37°C. After removal of the supernatant and two washing steps with PBS adherent cells were treated with different concentrations of rhamnolipids or derivatives for additional 24 h and the biofilms were quantified using CV as described above. Biofilm dispersion assays were performed in six wells and repeated three times.

MRC5 human lung fibroblasts were obtained from the ATCC. Cells were maintained as monolayer cultures in RPMI-1640 supplemented with 100 mg L^-1 streptomycin, 100 U mL^-1 penicillin and 10% (v/v) FBS (all from Sigma, Munich, Germany). Cells were grown in a humidified atmosphere of 95% air and 5% CO₂ at 37°C.

Cytotoxicity on MRC5 cells was evaluated with 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT). The assay was carried out after 48 h of cell incubation in the media containing test compounds at different concentrations and the viability was measured as described (Mihajlovic et al., 2012). The results are presented as the percentage of the control (untreated cells) that was arbitrarily set to 100%.

The results were analyzed by Student’s t-test using SPSS version 20 software. A P-value lower than 0.05 was considered as statistically significant.

The rhizosphere of medicinal plant G. hederacea L. (ground ivy) proved a prolific source of aerobic bacteria with 32 morphologically differing isolates collected from this source (data not shown). Culture extracts from these isolates were screened for their ability to inhibit biofilm formation in P. aeruginosa PAO1 using CV based assay in 96-well polystyrene plates. During the preliminary screening, the crude cell extract of the isolate BV152.1 showed the most potent ability to inhibit 60% of biofilm formation when applied in the concentration of 500 μg mL^-1 while no bactericidal activity of the extract was observed. This isolate was selected for structural characterization of its secondary metabolites as the most promising one. The isolate BV152.1 was identified as a member of the Gram-positive genus Lysinibacillus on the basis of 16S rRNA gene sequence analysis. Its 16S rRNA gene sequence (GenBank access No. KY933395) showed 99% similarity with type strains of Lysinibacillus louembei NM73 (NR_145586.1), L. meyeri WS 4626 (NR_117577.1), and L. odysseyi YH2 (KM873372.1).

In order to identify the component with biofilm formation inhibitory activity produced by Lysinibacillus sp. BV152.1, 1 L culture grown for 7 days at 30°C was extracted with ethyl acetate and the total mass of 800 mg crude cell extract was obtained. The crude cell extract was subsequently fractionated, the activity of each fraction measured and the fraction designated as F3 was identified as the most active one. After further purification, the total mass of F3 was determined to be 60 mg and it showed 50% inhibition of biofilm formation when applied in a concentration of 50 μg mL^-1 (Table 1).

¹H-NMR and ¹³C-NMR spectra of F3 showed the presence of fatty acid chains and carbohydrate moieties (Figure 1A) indicative of the characteristic spectra of bacterial di-rhamnolipids. The peak observed at δ 0.88 ppm indicated the presence of terminal methyl group and the peak at δ 1.26, characteristic for methylene group, indicated the presence of a straight-chain fatty acid. The peaks observed at δ 4.15–3.36 and peaks for two anomeric protons at δ 4.90 confirmed the presence of two carbohydrate units. The ¹³C-NMR analysis demonstrated two carbonyl peaks at δ 173.7 and 171.4, two anomeric carbon peaks at δ 102.5 and 94.5 and expected number of peaks for carbohydrate and fatty acid units. In general, the spectral characteristics obtained for F3 (provided in Supplementary material) were in good agreement with the data published in the literature for di-rhamnolipids (Sharma et al., 2007).

The exact lengths of the fatty acid chains were deduced from HPLC-MS data (Figure 1B). Sodiated molecular ions, [M + Na]+, in the ESI+MS data were observed at m/z 701 (Rha-Rha-C10-C12, 11.16 min, 9%), 699 (Rha-Rha-C10-C12:1, 10.68 min, 8%), 673 (Rha-Rha-C10-C10, 10.03 min, 63%, major), and 645 (Rha-Rha-C8-C10, 8.96 min, 17%), 527 (Rha-C10-C10, 10.86 min, 3%).

Considering the identified structural characteristics of F3, we have selected commercially available P. aeruginosa sp. derived rhamnolipids (R90), as the model and the control compound. R90 was determined to contain mono- vs. di-rhamnolipid form in 4:1 ratio (AGAE Technologies, Figure 1C) with the major component of Rha-Rha-C10-C10 in more than 90%.

Di-rhamnolipids from Lysinibacillus sp. BV152.1 (F3) containing mainly di-rhamnolipids with Rha-Rha-C10-C10 63% showed dose-dependent anti-biofilm formation activity in P. aeruginosa PAO1 with 50 μg mL^-1 determined as BFIC₅₀ (Figure 2A). In order to assess whether di-rhamnolipids from Lysinibacillus sp. BV152.1 affected biofilm growth, P. aeruginosa PAO1 cells were left to attach to the surface of microtiter wells for 2 h prior to incubation with F3 for 24 h. Inhibition of biofilm formation was similar in the presence or absence of the cell adhesion phase showing that di-rhamnolipids affected both cell attachment and biofilm growth (Figure 2A).

Anti-biofilm properties of di-rhamnolipids from Lysinibacillus sp. BV152 were compared to that of commercially available rhamnolipids obtained by fermentation of P. aeruginosa sp. (R90) (Figure 2B). Di-rhamnolipids (di-Rha) from this sample were also purified and used for the comparison. Rhamnolipids mixture (R90) and purified di-Rha from P. aeruginosa showed slightly lower activity against P. aeruginosa PAO1 biofilm formation (BFIC₅₀ = 75 μg mL^-1) comparing to F3. The effects of R90 and di-Rha on biofilm formation were comparable when the treatments were applied without cell adhesion phase. However, when bacteria were left to attach to the surface prior to treatments, di-Rha stimulated biofilm formation when added in concentrations up to 50 μg mL^-1. Di-rhamnolipids from both sources showed an equal potency in inhibition of biofilm formation at concentrations above 50 μg mL^-1.

Di-rhamnolipids from Lysinibacillus sp. BV152.1 efficiently inhibited bacterial adhesion to the polystyrene surface of microtiter plates and also to the surface of silicone catheters or glass coverslips as visualized by SEM and fluorescent microscopy, respectively (Figure 3).

Production of rhamnolipids in P. aeruginosa enables its swarming motility (Caiazza et al., 2005). Therefore, the effect of di-rhamnolipids on P. aeruginosa PAO1 swarming motility was analyzed next and the results showed that both F3 and R90 stimulated swarming in dose-dependent manner (Supplementary Figure S1). The observed effect was more prominent in the presence of di-rhamnolipids from Lysinibacillus sp. BV152.1.

The semi-synthetic approach of generating amide derivatives from two different sources of di-Rha was straightforward, with products obtained in high purity and yields from 25 to 55% (Figure 4). Products were characterized using NMR and LC-MS (Supplementary Figures S2–S5). In addition, silylated derivative (di-Rha-TBDMS) was also generated for the control purposes. Physico-chemical parameters for all compounds were calculated revealing the HLB value ranging from 8.72 to 11.09 for di-Rha and amide derivatives, while this value for di-Rha-TBDMS was 5.35 (Figure 4). It is known that compounds with HLB 7–11 are considered wetting agents and water in oil emulsifiers, while the ones having HLB between 4 and 6 as water in oil emulsifiers (Pasquali et al., 2008; Muller et al., 2012). pKa was three times higher for amide derivatives in comparison to di-Rha and di-Rha-TBDMS indicating that they were weaker acids. logP was between 3.36 and 5.07 for di-Rha and amide derivatives, while this value for di-Rha-TBDMS was 3–5 times higher, suggesting higher hydrophilicity of amide derivatives (Figure 4). Spectral data for the new compounds are provided in Supplementary material.

Antibacterial activity of all derivatives obtained from both rhamnolipids sources against P. aeruginosa PAO1, P. aeruginosa DM50, S. aureus ATCC 25923, S. aureus MRSA and S. marcescens was addressed. Neither F3 nor di-Rha derivatives obtained from Lysinibacillus sp. BV152.1 exhibited antibacterial activity at concentrations up to 500 μg mL^-1 against any of the bacterial species tested (Table 1). None of the derivatives synthetized from P. aeruginosa di-Rha affected P. aeruginosa and S. marcescens growth. However, P. aeruginosa derivatives di-Rha-Pip and di-Rha-Mor exhibited bactericidal activity against S. aureus ATCC 25923 strain and S. aureus MRSA with MIC concentrations 62.5 μg mL^-1. Derivative Rha-Bn from P. aeruginosa demonstrated the same antibacterial activity against S. aureus MRSA.

Cytotoxicity of rhamnolipids mixture (R90), di-Rha mixture (F3), purified di-Rha and their derivatives was analyzed (Figure 5). Viability of human lung fibroblasts (MRC5) was not affected in the presence of F3, di-Rha and di-Rha-TBDMS when applied in concentrations up to 100 μg mL^-1. Rhamnolipids mixture R90 showed cytotoxic effect with IC₅₀ value 50 μg mL^-1. Derivatization of di-Rha substantially increased their cytotoxicity reaching almost 100% cells killing at concentrations of 25 μg mL^-1. These concentrations were 2.5-fold lower than their MIC concentration values against S. aureus strains (Table 1) and potentially are limiting factor in their further development as antibiotics.

Anti-biofilm properties of di-rhamnolipids and their derivatives were examined by addressing their influence on biofilm formation, cell adhesion and disruption of pre-formed biofilms. Biofilm formation in P. aeruginosa PAO1 was quantified in the presence or absence of di-rhamnolipids and their derivatives. Derivative di-Rha-Mor from Lysinibacillus sp. BV152.1 showed increased anti-biofilm formation activity compared to di-rhamnolipids mixture F3 (BFIC₅₀ = 12.5 μg mL^-1) inhibiting 90% biofilm formation at 100 μg mL^-1 (Figures 6A, 7). Anti-biofilm formation activity of di-Rha-Bn and di-Rha-Pip was twofold lower than that of F3 (BFIC₅₀ = 100 μg mL^-1). Amide derivatization of di-Rha from P. aeruginosa significantly improved activities of all derivatives with BFIC₅₀ = 10 μg mL^-1 (Figures 6B, 7). However, maximum activity was reached already at 50 μg mL^-1 with biofilm formation inhibition up to 60%. Derivatives di-Rha-TBDMS from both sources, showed no anti-biofilm formation activity.

To measure anti-adhesion activity of di-rhamnolipids and their amide derivatives, GFP fluorescence of P. aeruginosa PAO1-GFP was used to quantify bacterial adhesion 2 h after inoculation. Relative to the amount of adhered bacteria without treatment (containing 0.1% DMSO), derivatives di-Rha-Bn and di-Rha-Pip from Lysinibacillus sp. BV152.1 inhibited PAO1-GFP adhesion by 50% at concentrations 10 μg mL^-1, while di-Rha-Mor showed 50% adhesion inhibition at 50 μg mL^-1 (Figure 6C). Again, derivatives from P. aeruginosa exhibited less prominent anti-adhesive activities inhibiting P. aeruginosa PAO1-GFP adhesion up to 35% at concentrations 100 μg mL^-1 (Figure 6D). Derivative di-Rha-TBDMS showed no significant anti-adhesion activity. Importantly, di-Rha and their derivatives did not affect GFP expression nor quenched its fluorescence (Supplementary Figure S10).

Potential to disperse formed biofilm is often more relevant to medical applications, and more challenging than inhibition of biofilm formation. Therefore, the ability of di-rhamnolipids and their amide derivatives to disperse 24 h old P. aeruginosa biofilms was examined next (Figures 6E,F, 8). Di-rhamnolipids from Lysinibacillus sp. BV152.1 (F3) were more effective in biofilm disruption with BDIC₅₀ 10 μg mL^-1 (BDIC₅₀ is the concentration of compound that caused 50% biofilm disruption) comparing to di-Rha from P. aeruginosa which disrupted biofilms up to 20% at concentrations 100 μg mL^-1. The strongest biofilm dispersion activity was measured for di-Rha-Mor derivatives from both sources, with BDIC₅₀ values 12.5 and 50 μg mL^-1 from Lysinibacillus sp. BV152.1 and P. aeruginosa, respectively. Both di-Rha-Mor derivatives were able to disrupt more than 80% of pre-formed biofilms when applied at 100 μg mL^-1.

Taken together, these results demonstrated that amide derivatization improved di-rhamnolipids anti-biofilm properties, with di-Rha-Mor derivative being the most active compound. The results were confirmed by comparing anti-biofilm formation activities of di-rhamnolipids and their amide derivatives against different bacterial species (Tables 2, 3). The improvement of rhamnolipids anti-biofilm formation activity with amide derivatization was more prominent with di-rhamnolipids from Lysinibacillus sp. BV152.1. Anti-biofilm formation activity of these derivatives was species specific with the highest activity observed for di-Rha-Bn and di-Rha-Pip against S. aureus ATCC 25923 (>90% inhibition). All three derivatives were active against S. marcescens biofilm formation (>80%), while the highest activity against P. aeruginosa biofilms was observed for di-Rha-Mor inhibiting 80% biofilm formation. The most active P. aeruginosa di-Rha derivative was also di-Rha-Mor, with improved anti-biofilm formation activity compared to di-Rha against S. aureus and S. marcescens (70% and 85% inhibition at 50 μg mL^-1, respectively). The strongest inhibition of P. aeruginosa PAO1 biofilm formation was achieved with P. aeruginosa di-Rha-Bn and di-Rha-Pip treatments, while none of the derivatives affected biofilm formation in clinical isolate P. aeruginosa DM50 (Table 2). Derivatization of di-rhamnolipids from both sources also improved their biofilm dispersion activity against P. aeruginosa DM50, S. aureus ATCC 25923 and S. aureus MRSA (Table 3). Dispersion of S. marcescens biofilms was efficient only with di-Rha-Bn and di-Rha-Mor from Lysinibacillus sp. BV152.1 (63% and 58% biofilm dispersion at 50 μg mL^-1, respectively). Di-Rha from both sources and the other derivatives showed no biofilm dispersion activity against this bacterium, most likely be due to slightly different carbon chain length between di-rhamnolipid sources.