Characterization of Potential Virulence, Resistance to Antibiotics and Heavy Metals, and Biofilm-Forming Capabilities of Soil Lignocellulolytic Bacteria

The soil microbiota is essential in cycling carbon, nitrogen, and other nutrients. Soil bacteria are biotransformation agents of soil matter and nutrients widely used in industrial processes. The interactions of soil bacteria are very complex. Changes in diversity within one trophic group or functional guild can alter the diversity, prevalence, and functioning, leading to the development of various strategies for their survival [Ning et al., 2021].

While soil bacteria develop strategies to better survive in their environment, they can become more harmful to humans in terms of infection development and lead to higher medical costs, prolonged hospital stays, and increased mortality rates [Mogrovejo et al., 2020]. Various reports revealed that soil bacteria developing virulence factors are increasing annually worldwide due to the contamination with high concentrations of toxic metals from agrochemicals, industrial wastewater, and gas and coal mining [Yang et al., 2022]. In such circumstances, they exhibit their pathogenicity through several mechanisms such as (1) the contribution of pili/flagella/fimbriae/adhesins to adherence, autoaggregation, biotic and abiotic surface colonization; (2) the role of outer membrane lipopolysaccharide (LPS) in biofilm formation, resistance to antibiotics and heavy metals, and complement-mediated cell killing; (3) the role of diffusible signal factor in quorum sensing that mediates extracellular enzyme production, LPS synthesis, microcolony formation, antibiotic and heavy metal tolerance; and (4) extracellular enzyme production such as protease, esterase, DNase, RNase, lipase, hemolysin, gelatinase, and fibrinolysin [Abbott et al., 2011; Brooke, 2012; Kalidasan et al., 2018; Odeyemi and Sani, 2019; Rhen, 2019].

Six cellulolytic bacteria, Paenarthrobacter sp., Hymenobacter sp., Mycobacterium sp., Stenotrophomonas sp., Chryseobacterium sp., and Bacillus sp., isolated from isolated soil samples collected in Kingfisher Lake and the University of Manitoba campus showed promising activities in the fermentation process [Mokale et al., 2022a, 2022b]. However, their innocuity has not been investigated yet, a primary characteristic in lignocellulosic biomass bioprocess industries. Several virulence factors were reported in Mycobacterium [Ly and Liu, 2020], Stenotrophomonas [Kalidasan et al., 2018], Chryseobacterium [Mwanza et al., 2022], and Bacillus [Mbhele et al., 2021] species collected from various sources. Therefore, this study investigated potential virulence factors, antibiotic and heavy metal resistance, solvent adhesion, and biofilm-forming capabilities of these soil cellulolytic bacteria.

Six bacterial isolates with cellulolytic properties (cellulase and glucose isomerase activities) were characterized using soil samples from Kingfisher Lake and the University of Manitoba campus. These isolates were identified as Paenarthrobacter sp. MKAL1, Hymenobacter sp. MKAL2, Mycobacterium sp. MKAL3, Stenotrophomonas sp. MKAL4, Chryseobacterium sp. MKAL5, and Bacillus sp. MKAL6 with the NCBI accession numbers ON442553, ON442554, ON442555, ON442556, ON442557, and ON442558, respectively [Mokale et al., 2022a, 2022b]. Bacteria were maintained in Tryptic soy broth (TSB) at 30°C for subsequent tests.

Hemolytic, proteolytic, and lipase activities and capsule production were screened using blood agar, skim milk agar, tween 80 agar, and Congo red agar, respectively [Lamari et al., 2018]. Overnight bacterial cultures were inoculated in sterile Petri dishes containing respective culture medium and then incubated at 30°C for 48 h. After incubation, a clear zone of hydrolysis around the bacterial isolate indicated the presence of proteolytic and lipase activities. A greenish-gray or brownish discoloration around the colony revealed α-hemolysis (α-hemolysin production), while the clear zone appearance around the colony showed β-hemolysis (β-hemolysin production). The absence of coloration changes or zone appearance indicated γ-hemolysin production (no cell blood lysis). Black colonies were capsule producers, while red colonies were non-capsule producers.

Autoaggregation assay was performed according to Escamilla-Montes et al. [2015]. Overnight bacterial cultures were collected, centrifuged at 5,000 × g for 15 min, washed twice, and suspended in phosphate-buffered saline (PBS, pH 7.4). Cell density was adjusted to the optical density of 0.55–0.60 at 600 nm (A₀). Bacterial cell suspensions (4 mL) were mixed by vortexing for 10 s and incubated at room temperature for 24 h. After 3, 6, and 24 h, 0.1 mL of the upper suspension was transferred to another 3.9 mL of PBS, and absorbance was measured at 600 nm. PBS was used as a blank.

$Autoaggregation%=1−At/A0×100$⁠, where A_t represented the absorbance at time t = 3, 6, or 24 h and A₀, the absorbance at t = 0.

The bacterial adherence to hydrocarbons (BATH) test was used to evaluate the bacterial hydrophobicity potential [Borghi et al., 2011]. This test analyzed microbial linkage to n-octane (apolar solvent), chloroform (polar acid solvent), and ethyl acetate (basic polar solvent). Bacterial strains were grown in TSB at 30°C for 24 h. The bacterial culture was centrifuged at 5,000 × g for 15 min, the supernatant was discarded, and pellets were washed twice with PBS (pH 7.4). The density of cells was adjusted to the optical density of 0.55–0.60 at 600 nm (A₀). The test mixture was composed of cell suspension (4 mL) and 1 mL of n-octane, chloroform, and ethyl acetate in individual glass tubes and then vortexed for 1 min. The mixture was decanted into two phases at room temperature for 30 min. The supernatant was discarded, and absorbance was read at 600 nm (A₁). Hydrophobic ability was estimated according to the formula:

$Hydrophobicity%=A0−A1/A0×100$⁠, and isolate is classified into three categories: not hydrophobic (<20%), moderate (20–50%), and strong (>50%).

Resistance of the isolates to heavy metals (Co²⁺, Cd²⁺, Cr³⁺, Zn²⁺, Hg²⁺, Cu²⁺, Mn²⁺, Ni²⁺, Ba²⁺, and Pb²⁺) was carried out by inoculating overnight bacterial culture on Tryptic soy agar Petri dishes containing various concentrations of metal (50, 150, 300, 450, 600, and 750 µg/mL) [Marzan et al., 2017]. The metal was loaded into the medium in the form of chloride salt, and the pH of the medium was adjusted to 7 by sodium hydroxide or hydrochloride if necessary. Visible growth of isolates was observed for 24 and 48 h at 30°C. Minimum inhibitory concentration (MIC) was the lowest concentration inhibiting bacterial growth.

The biofilm-forming capacity of bacterial strains was carried out by adhesion to polystyrene [Chaieb et al., 2011]. Isolates were grown in TSB at 30°C and then diluted to 1:100 w/v (in TSB with 2% glucose). Aliquots of cell suspensions (200 µL) were transferred to 96-well microtiter plates and incubated at 35°C for 24 h. Plates were washed twice with PBS and dried. The well with sterile TSB alone was used as a control. Adherent strains were fixed with ethanol (95%) and stained with 100 µL crystal violet (CV) (1% w/v) solution for 5 min. Microplates were washed and air-dried. Biofilm-forming ability was measured at 570 nm. The experiment was done in triplicate. Biofilm formation was interpreted as follows: highly positive (OD₅₇₀ ≥ 1), moderately to weakly positive (0.1 ≤ OD₅₇₀ < 1), or negative (OD₅₇₀ ≤ 0.1).

The bacterial susceptibility to antibiotics was investigated by determining the diameter of inhibition zones (DIs) and MICs using the agar disc diffusion and microdilution methods [CLSI, 2008]. Bacterial cell suspensions were prepared at 1.5 × 10⁸ colony-forming units per mL (CFU/mL) corresponding to the McFarland 0.5 turbidity standard. The bacterial suspension (100 μL) was spread in Petri dishes containing sterile MHA (20 mL). Antibiotic discs were dropped on the surface of MHA dishes and diffused for 15 min before incubation at 35°C for 24 h. Antibiotics tested were ampicillin (10 μg), novobiocin (30 μg), bacitracin, tetracycline (30 μg), erythromycin (15 μg), chloramphenicol (30 μg), penicillin (10 units), hygromycin B (50 μg), lincomycin (15 μg), phleomycin (50 μg), kanamycin (30 μg), trimethoprim (15 μg), and ciprofloxacin (15 μg). Dishes without antibiotics were used as blank.

For the microdilution method, a two-fold dilution of antibiotics (v/v medium, inoculum, and water-soluble antibiotics) and negative control (v/v medium and inoculum) were included. Each well of a 96-well sterile microtiter plate received Mueller-Hinton broth (100 μL), antibiotic (100 μL), and bacterial inoculum (1.5 × 10⁸ CFU/mL), and plates were covered and incubated at 35°C for 24 h. After incubation, 50 μL of aqueous p-iodonitrotetrazolium violet (bacterial growth indicator) was added to the wells and incubated for 30 min. The MIC value was the lowest concentration of antibiotics that completely inhibited cell growth (when the solution remained clear in the well after incubation with p-iodonitrotetrazolium violet). Ampicillin, chloramphenicol, ciprofloxacin, trimethoprim, kanamycin, and lincomycin were used at concentrations ranging from 128 to 1 µg/mL. The antibiotic with the highest inhibitory effect (lowest MIC values) was used to explore its action on bacterial sensibility.

The antimicrobial efficacy testing was performed to evaluate the inhibitory effect of ciprofloxacin over time [Tsuji et al., 2008]. Bacterial isolates were subcultured and diluted to obtain an inoculum size of 5 × 10⁶ CFU/mL. Concentrations of ciprofloxacin equal to MIC, 2MIC, and 4MIC were prepared and transferred into test tubes containing Mueller-Hinton broth and bacterial inoculum. Tubes were incubated at 35°C. Aliquots of medium (1 mL) were collected at 0, 2, 4, 6, 12, and 24 h, inoculated aseptically into MHA Petri dishes, and incubated at 35°C for 24 h. A control test was carried out without the antibiotic. The number of viable organisms was counted as CFU. Graphs of the log CFU/mL were plotted against time.

The effect of ciprofloxacin on cell membrane integrity and membrane permeability was conducted using the protocol described by Devi et al. [2010]. Overnight bacterial cultures were centrifuged (5,000 × g, 15 min), and the supernatant was discarded. Pellets were washed twice using sterile distilled water and then suspended in PBS (pH 7.4) to obtain a cell density of 0.55–0.60 at 600 nm. Cell suspensions were treated with ciprofloxacin (MIC and 4MIC). Cell suspensions without antibiotic treatment were used as negative controls. For effect on membrane integrity, samples were incubated at 35°C for 1 h under agitation (200 rpm), then centrifuged (12,000 × g, 15 min), and supernatants were read at 260 nm. Recordings were expressed as percentages of the extracellular UV-absorbing materials released by cells.

All experiments were performed in triplicate. Data were expressed as mean ± SD. Statistical analysis was carried out using one-way analysis of variance (ANOVA) followed by Student-Newman-Keuls multiple comparison tests using GraphPad Prism 5 Windows software. Differences between values were considered significant at p < 0.05.

Among the six bacterial isolates studied, only Bacillus sp. produced capsules. Hymenobacter sp., Mycobacterium sp., Chryseobacterium sp., and Bacillus sp. exhibited proteolytic activities. Hymenobacter sp., Chryseobacterium sp., and Bacillus sp. showed lipase activity. No hemolytic properties were observed (Table 1).

The autoaggregation ability of bacterial strains was investigated based on their sedimentation features. Their autoaggregation capability increased with incubation time (Fig. 1). The aggregation potential increased from 31.09 to 56.36%, 26.18 to 59.95%, 33.30 to 58.87%, 21.11 to 53.16%, 18.13 to 46.30%, and 29.11 to 55.11% for Bacillus sp. MKAL6, Hymenobacter sp. MKAL2, Chryseobacterium sp. MKAL5, Paenarthrobacter sp. MKAL1, Mycobacterium sp. MKAL3, and Stenotrophomonas sp. MKAL4, respectively. Hymenobacter sp. MKAL2 showed the highest autoaggregation potential (59.95%) at 24 h, while Mycobacterium sp. MKAL3 showed the lowest ability (46.30%).

Cell surface hydrophobicity of bacterial isolates analyzed microbial linkage to n-octane (apolar solvent), chloroform (polar acid solvent), and ethyl acetate (basic polar solvent). Hydrophobic cell surface showed adherence to n-octane (6.66–31.08%), chloroform (19.88–45.58%), and ethyl acetate (8.57–58.16%) (Fig. 2). The hydrophobicity ability of Bacillus sp. MKAL6 increased with solvent polarity, while that of Mycobacterium sp. MKAL3 decreased with solvent polarity. All isolates showed a moderate hydrophobicity to chloroform except for Hymenobacter sp. MKAL2 (19.88%). Paenarthrobacter sp. MKAL1 (29.76%), Stenotrophomonas sp. MKAL4 (43.28%), and Chryseobacterium sp. MKAL5 (37.31%) exhibited the highest hydrophobicity ability to chloroform, while Hymenobacter sp. MKAL2 (21.77%) and Mycobacterium sp. MKAL3 (31.08%) showed the highest adhesion effect to n-octane. However, Bacillus sp. MKAL6, a capsule producer, exhibited a strong hydrophobicity for ethyl acetate (58.16%).

The biofilm-forming capacity of isolates was estimated by adhesion to polystyrene. Paenarthrobacter sp. MKAL1 (OD₅₇₀ = 2.985), Hymenobacter sp. MKAL2 (OD₅₇₀ = 2.994), Mycobacterium sp. MKAL3 (OD₅₇₀ = 2.000), Stenotrophomonas sp. MKAL4 (OD₅₇₀ = 1.286), and Bacillus sp. MKAL6 (OD₅₇₀ = 3.295) showed strong biofilm-forming capacity (OD₅₇₀ ≥ 1), while Chryseobacterium sp. MKAL5 (OD₅₇₀ = 0.601) exhibited moderate ability (0.1 ≤ OD₅₇₀ < 1) (Fig. 3).

Resistance of isolates to heavy metals (Co²⁺, Cd²⁺, Cr³⁺, Zn²⁺, Hg²⁺, Cu²⁺, Mn²⁺, Ni²⁺, and Pb²⁺) were carried out at the concentrations of 50, 150, 300, 450, 600, and 750 µg/mL. MICs of heavy metals were determined. All isolates showed variable degrees of resistance to heavy metals (Table 2). Cadmium inhibited bacterial growth at the lowest concentration tested (MIC = 50 µg/mL). Except for Stenotrophomonas sp. MKAL4 (MIC = 50 µg/mL), cobalt inhibited cell growth at the highest concentration tested (MIC = 750 µg/mL). Copper and mercury inhibited the growth of all isolates tested with MICs ranging from 50 to 450 µg/mL. Chromium, lead, zinc, nickel, and manganese did not inhibit bacterial growth except for Stenotrophomonas sp. MKAL4 at 600 µg/mL.

The bacterial susceptibility test was performed using various antibiotics (Fig. 4). In the agar-well diffusion method, all isolates were resistant to lincomycin with DIs ranging from 0.0 to 18.6 mm. Hymenobacter sp. was resistant to novobiocin (DI = 9.6 mm), bacitracin (DI = 0.0 mm), and tetracycline (DI = 10.1 mm), while Mycobacterium sp. (DI = 8.3 mm) was intermediate to bacitracin (Table 3). In the microdilution method, all antibiotics exerted an inhibitory effect against bacteria isolates with MIC ranging from 0.25 to 512 µg/mL (Table 4). Ciprofloxacin exhibited the highest inhibitory activity (MIC = 0.25–0.5 µg/mL), while kanamycin exhibited the lowest activity (MIC = 4–512 µg/mL). Paenarthrobacter sp. (MIC = 0.25–128 µg/mL), Mycobacterium sp. (MIC = 0.25–256 µg/mL), Hymenobacter sp. (MIC = 0.25–512 µg/mL), Stenotrophomonas sp. (MIC = 0.5–512 µg/mL), Chryseobacterium sp. (MIC = 0.5–512 µg/mL), and Bacillus sp. (MIC = 0.5–256 µg/mL) exhibited variable susceptibilities to antibiotics. As ciprofloxacin showed the highest inhibitory effect (lowest MIC values), its action was investigated on bacterial susceptibility over time, cell membrane, and biofilm formation.

Ciprofloxacin was tested for antibacterial efficacy over time (24 h) at different concentrations (MIC, 2MIC, and 4MIC). The time-kill kinetics profile of ciprofloxacin against bacterial isolates showed a reduction in viable cell number over 24 h compared to the control (non-treated cells). The viable cell number decreased with increasing antibiotic concentrations. Ciprofloxacin exhibited the highest inhibitory effect against Chryseobacterium sp. MKAL5 and Bacillus sp. MKAL6 at the concentration of 4MIC. The overall effect of ciprofloxacin was bactericidal at all tested concentrations (Fig. 5).

The effect of ciprofloxacin on the bacterial cell membrane was performed by quantifying the release of UV-absorbing materials (OD₂₆₀), an index of membrane integrity damage and loss. The results are presented in Figure 6a. After treatment with ciprofloxacin at MIC and 4MIC, a slight increase in OD was observed in Bacillus sp. (0.139–0.176 and 0.139–0.185), Chryseobacterium sp. (0.140–0.162 and 0.140–0.169), Paenarthrobacter sp. (0.144–0.174 and 0.144–0.177), and Mycobacterium sp. (0.140–0.165 and 0.140–0.184). However, no change in OD was observed in Hymenobacter sp. MKAL2 and Stenotrophomonas sp. MKAL4. The action of ciprofloxacin on bacterial cell membrane permeability is shown in Figure 6b. CV uptake significantly (p < 0.001) increased after treatment with ciprofloxacin at MIC and 4MIC in Stenotrophomonas sp. MKAL4 (62.06 and 62.87%), Chryseobacterium sp. MKAL5 (41.90 and 46.56%), and Bacillus sp. MKAL6 (50.43 and 52.89%) compared to untreated cells (23.87, 32.56, and 52.35%, respectively).

Ciprofloxacin was tested for biofilm formation inhibition capability at different concentrations (8MIC–1/16MIC). Ciprofloxacin significantly (p < 0.001) inhibited the biofilm formation in Bacillus sp. (69.27–39.58%), Hymenobacter sp. (76.13–55.13%), Chryseobacterium sp. (96.98–75.96%), Paenarthrobacter sp. (71.90–38.75%), Mycobacterium sp. (80.07–60.38%), and Stenotrophomonas sp. (86.54–68.03%) (Fig. 7).

Soil bacteria adhere to diverse surfaces and promote biofilm formation in a protective and self-produced matrix responsible for disease emergence and resurgence [Ning et al., 2021]. Thus, potential virulence factors, antibiotic and heavy metal resistance, solvent adhesion, and biofilm-forming capabilities in these strains were investigated.

Various enzymes such as hemolysins [Elliott et al., 1998], proteases [Ingmer and Brøndsted, 2009], and lipases [Jaeger et al., 1994] have been implicated in microbial virulence. In our study, Hymenobacter sp. MKAL2, Mycobacterium sp. MKAL3, Chryseobacterium sp. MKAL5, and Bacillus sp. MKAL6 exhibited proteolytic activities. Hymenobacter sp. MKAL2, Chryseobacterium sp. MKAL5, and Bacillus sp. MKAL6 showed lipase activity. No hemolytic properties were observed. The hemolytic activities of Bacillus [Dabiré et al., 2022], Chryseobacterium [Sud et al., 2020], Mycobacterium [Augenstreich et al., 2020], and Stenotrophomonas genera [Peters et al., 2020] were reported. However, only Bacillus sp. MKAL6 produced capsules. Although capsular structures can be adhesins for some bacteria, most capsules in pathogenesis protect bacteria against host immune mechanisms. The capsule production in Bacillus cereus and Bacillus anthracis is required for their virulence [Beesley et al., 2010; Baldwin, 2020].

Adherence is the first step in bacterial pathogenesis. It is related to the capacity for autoaggregation, hydrophobicity, and biofilm formation of bacterial cells. In autoaggregation, bacteria (same type) form multicellular clumps, generally mediated by self-recognizing surface structures such as capsules and proteins, leading to biofilm formation. Moreover, cell surface hydrophobicity increases microbial cell propensity to adhere to surfaces [Nwagu et al., 2020]. All strains showed high autoaggregation properties (18.13–59.95%) [Nwagu et al., 2020]. reported about 53.37% of autoaggregation in Bacillus cereus KY746353.1 isolated from Parkia biglobosa (traditional fermented African locust bean seeds), which was similar to the potential observed in Bacillus sp. MKAL6 (56.36%). However, the autoaggregation of Bacillus subtilis P223 isolated from Kimchi (Korean food) was 93.42% after 24 h of incubation [Jeon et al., 2017]. Manhar et al. [2015] reported that the highest autoaggregation potential of Bacillus amyloliquefaciens AMS1 isolated from traditional fermented soybean (Churpi) was 75.5% after 24 h [Benladghem et al., 2020]. revealed that Stenotrophomonas maltophilia could form cellular aggregates of 26.13% after 24 h, which was lower than that of Stenotrophomonas sp. MKAL4 (55.11%). The cell hydrophobicity varied with the isolates tested. The differences in affinity may be due to the capsular material, appendages on the cell surface or composition, and content of different LPSs [Ning et al., 2021]. Bacillus sp. MKAL6 exhibited a strong hydrophobicity for ethyl acetate (58.16%) and moderate hydrophobicity for chloroform. This indicates that this bacterial strain had an affinity for electron acceptance (ethyl acetate) and electron donation (chloroform). The hydrophobicity capabilities of Bacillus sp. in this study were comparatively lower than those reported by Kuebutornye et al. [2020]. Also, Amenyogbe et al. [2021] revealed the adhesion of Bacillus sp. RCS1 (97.2%) and Bacillus cereus (97.1%) isolated from Cobia fish (Rachycentron canadum) to xylene, chloroform, and ethyl acetate were strong. Paenarthrobacter sp. MKAL1, Hymenobacter sp. MKAL2, Mycobacterium sp. MKAL3, Stenotrophomonas sp. MKAL4, and Bacillus sp. MKAL6 showed strong biofilm-forming capacity, while Chryseobacterium sp. MKAL5 displayed moderate ability. Many investigators revealed the biofilm-forming ability of these bacteria species [Esteban and García-Coca, 2018; Kim et al., 2020; Bostanghadiri et al., 2021; Sornchuer et al., 2022]. Bacillus sp. MKAL6, which exhibited higher hydrophobic ability, adhered more to polystyrene. It was reported that bacteria with high hydrophobic surface affinity displayed a high biofilm-forming ability [Ning et al., 2021].

The autoaggregation and biofilm formation confer antimicrobial and heavy metal resistance, contributing to the metabolic cooperation, production of virulence factors, survival, and persistence in some niches or hosts. Also, multiple antibiotic resistances are associated with resistance/tolerance to heavy metals in soil bacteria. For different genera, the genes for heavy metal resistance are often plasmid encoded. Since these genes are clustered on the same plasmids, heavy metals and antibiotics are environmental factors which exert selective pressure on the populations of these plasmid-harboring bacteria [Lodha et al., 2022]. Except for Stenotrophomonas sp. MKAL4, all strains tolerated a large amount of chromium, lead, zinc, nickel, and manganese. Many bacteria isolated from diverse sources, such as Bacillus sp. [Glibota et al., 2020; Nath et al., 2020; Alotaibi et al., 2021], Stenotrophomonas sp. [Agarwal et al., 2019; Nath et al., 2020], Chryseobacterium sp. [Glibota et al., 2020], Arthrobacter sp. [Pathak et al., 2020], and Mycobacterium sp. [Sepehri et al., 2017] were reported to tolerate high levels of heavy metals and have heavy metal resistance genes. All strains exhibited variable susceptibilities to antibiotics. Hymenobacter sp. MKAL2 was resistant to novobiocin, bacitracin, and tetracycline. Kang et al. [2018] showed that Hymenobacter defluvii, isolated from wastewater, was resistant to amikacin (30 μg), gentamicin (10 μg), and kanamycin (30 μg). Ciprofloxacin showed bactericidal effects against bacterial strains by reducing viable cell numbers over 24 h at all tested concentrations. However, Grillon et al. [2016] showed that ciprofloxacin exhibited a bacteriostatic effect on some Stenotrophomonas maltophilia up to 6 h, followed by a regrowth at 24 h. Ciprofloxacin slightly increased the release of UV-absorbing materials in Paenarthrobacter sp. MKAL1, Mycobacterium sp. MKAL3, Chryseobacterium sp. MKAL5, and Bacillus sp. MKAL6 at the concentrations of MIC and 4MIC. This suggests that ciprofloxacin weakly damaged the cytoplasmic membrane resulting in low leakage of bacterial isolates intracellular constituents (proteins and nucleic acids). However, no change in OD was observed in Hymenobacter sp. MKAL2 and Stenotrophomonas sp. MKAL4, indicating that ciprofloxacin did not affect the membrane integrity of these isolates. CV uptake significantly increased after treatment with ciprofloxacin at MIC and 4MIC in Stenotrophomonas sp. MKAL4, Chryseobacterium sp. MKAL5, and Bacillus sp. MKAL6. The increase in CV uptake might be attributed to the change in permeability and structure of the bacterial cell wall layer [Seukep et al., 2020]. Ciprofloxacin significantly inhibited biofilm formation in all bacterial strains. The potential of ciprofloxacin to reduce biofilm formation was reported [Kwiecińska-Piróg et al., 2013].

The findings of this study indicate that isolated bacteria species from soil exhibited some virulence factors. Hymenobacter sp. MKAL2, Chryseobacterium sp. MKAL5, and Bacillus sp. MKAL6 showed protease and lipase activities, while only Bacillus sp. MKAL6 produced capsules. All strains could aggregate, form biofilm, and adhere to organic solvents. All strains tolerated a high amount of chromium, lead, zinc, nickel, and manganese and were resistant to lincomycin. Although the phenotypic evaluation of virulence factors of bacteria can indicate their pathogenic nature, an in-depth genetic study of virulence, antibiotic and heavy metal resistance genes is required.

The authors sincerely thank all faculty members who directly or indirectly helped complete this research.

Ethical approval is not required for this study in accordance with local or national guidelines.

The authors have no conflict of interest to declare.

This project was supported by the Natural Science and Engineering Research Council of Canada (NSERC) Discovery Grant (RGPIN-2017-05366) to W.Q.

W.Q., C.C.X., Z.H.J., and A.L.M.K. contributed to the study conception and experimental design. A.L.M.K. carried out the material preparation and data collection, curation, and analysis. The manuscript draft was initially written by A.L.M.K. and edited by C.C., J.R.K., S.S., X.C., Y.Z., R.A.N.N., G.A.A., Z.H.J., C.C.X., and W.Q. W.Q. provided supervision and project administration. All authors have read and approved the final version of the manuscript.

The datasets generated during and analyzed during the current study are not publicly available for ethical reasons. Further inquiries can be directed to the corresponding author via email.