Isolation and Identification of S. aureus
During the study, isolates of S. aureus were recovered from different food commodities. For isolation and identification of S. aureus, the growth was monitored on differential and selective media. The isolates were streaked onto media such as Manitol Salt Agar (BioM), Staph-chromo agar (Merck), Staphylococcus 110 Agar (BioM), Baird-Parker Agar (Oxoid), DNase Agar (Merck), and Blood Agar (Oxoid). Following incubation, the growth on these selective media was examined for characteristic colony morphology indicative of S. aureus. Confirmation of S. aureus was performed using the Staph Latex kit (Prolix Latex Agglutination System) according to the manufacturer's instructions.
Biofilm Formation: S. aureus biofilms were developed using the previously described protocol (Ahmed et al., 2022). Briefly, S. aureus isolates were cultured in Tryptic Soy Broth (TSB) for 18 to 24 hours at 35°C. The bacterial cells were then separated by centrifugation at 10,000 rpm and washed with phosphate-buffered saline (PBS) at pH 7.4. The cell suspension was adjusted to a concentration corresponding to a 0.5 McFarland standard, resulting in approximately 1.5 × 108 CFU/mL. To study biofilm formation, TSB (pH 7.0) was prepared and distributed into 100 mL aliquots. Then, 100 µL of inoculum from a cell suspension with a concentration of 1.5 × 108 CFU/mL was added to each flask. A glass slide was placed in each flask to serve as a substrate for cell adhesion. The flasks were then incubated at 35°C for duration of 7 days without changing the media. During this incubation period, the conditions were kept constant to allow for the formation of biofilms, where microorganisms attach to the glass slides and develop into structured communities over the course of a week. To evaluate biofilm formation, the glass slides were carefully collected from the flasks and washed with PBS to remove unbound debris. The biofilms were fixed by microwaving for 30 seconds at 450 Watts and stained with a 0.1% crystal violet solution for 20 minutes. Subsequently, the slides were washed again with PBS to remove excess stain, and the biofilm-bound crystal violet was solubilized in 200 µL of an ethanol-acetone mixture (4:1 ratio). The optical density (OD) of each slide was measured at 578 nm using a spectrophotometer (UV/Visible spectrophotometer, Shimadzu Corporation, Kyoto, Japan).
Recovery of biofilm population
The biofilm positive slides underwent three washes with PBS (pH 7.0) to effectively remove debris and loosely bound cells from the biofilm consortia. This meticulous washing process aimed to eliminate any non-adherent or loosely attached material, ensuring that only cells firmly attached to the surface were retained for further analysis. By removing the non-specifically bound cells, the focus is placed on studying the biofilm's core structure and the cells that are firmly associated with it. Additionally, after the initial washes with PBS, the biofilm positive slides were subjected to further washing by vortexing for 30 seconds. This specific washing technique was employed to isolate and detach the cells that were directly attached to the glass surface. The vortexing action, facilitated the release of cells tightly adhered to the slide, allowing for their subsequent analysis or characterization (Mirani, Fatima, et al., 2018).
Population study
The cells, with a McFarland Standard of 0.5, were inoculated into 5 mL of TSB and incubated at 35°C in a shaking incubator set at 200 rpm. After 24 hrs, a 100 µL aliquot from this culture was taken and used to re-inoculate a fresh 5 mL of TSB. This procedure was repeated. Simultaneously, another aliquot was inoculated onto a Tryptone Soya agar (TSA) plate and incubated at 35°C for 96 hrs. The agar plates were examined every 24 hrs over the course of 96 hrs to monitor colony growth and development. The colonies were marked upon their initial appearance. Normal phenotypes were observed within 24 hours of incubation, displaying typical growth characteristics (diameter of > 1 mm, pigmented). In contrast, metabolically inactive cells, known as small colony variants, exhibited delayed growth and formed small, pinpointed non-pigmented transparent colonies. Typically, these small colony variants appeared after 48 hours of incubation (diameter of < 1 mm, pin-pointed, transparent and non-pigmented). In our study, all the cultures were maintained within the range of 1x105 to 1x106 CFU/mL, which indicates the density of viable cells in the cultures. To determine the viable counts, serial dilutions of the cultures were prepared in TSA, followed by overnight incubation at 35°C. This method allowed us to enumerate the viable bacterial colonies and estimate the number of colony-forming units (CFU) present in the original cultures (Ahmed et al., 2022; Mirani, Fatima, et al., 2018).
Colony Spreading Assay
To perform the modified colony spreading assay, a sterile petri dish was filled with 20 mL of TSB supplemented with 0.24% agar as described previously (Mirani, Khan, et al., 2018). After drying in a safety cabinet for 20 minutes, 2 µL (1.5 × 108) aliquots of 48-hour-old cell clumps with nanotubes and floating cells were separately spotted onto the center of the dried agar surface using a Micropipette (DragonLAB- United Kingdom). Following another 15 minutes of drying, the petri dish was incubated overnight at 35°C. The presence of giant colonies with branched arms was used as a positive indicator for colony spreading.
Autolysis
The Triton (T) X-100 induced autolysis was performed according to a previously published assay (Boyle-Vavra, Challapalli, & Daum, 2003). Briefly, the cells were harvested, washed twice with ice-cold water, and then re-suspended in the same volume of 0.05 M Tris-HCl (pH 7.2) containing 0.05% TX-100. Cells were incubated at 35°C with shaking for 30 minutes and checked for lysis by measuring the progressive decrease in absorbance (OD600). Autolysis was quantified as a percentage of the initial OD600 remaining at each sampling point.
Determination of Bacterial survival in TX-100
To perform the experiment, each culture suspension was diluted 10-fold in two different solutions: 0.1 mol/L PBS pH 7.0 (control) and TX-100. The aim was to achieve a final concentration of 0.5% w/v (8mM) of TX-100 for the treatment. The TX solution was prepared in 0.1 mol/L PBS pH 7.0. After dilution, all samples were incubated at 35°C for 30 minutes in a shaking incubator at 200RPM. Following incubation, the concentration of bacteria in each sample was determined. This was done by plating serial 10-fold dilutions of the samples in PBS (pH-7.0) and then pouring them onto TSA plates. Subsequently, all plates were incubated at 35°C for 48 hrs.
Scanning Electron Microscopy
Biofilm positive slides were divided into 4 mm sections and washed with distilled water to remove debris and negatively stained with 2% uranyl acetate for 30 s. Dehydration was carried out with ethanol (absolute) initially in 50% for 30 min, followed by 75% for 30 min and 95% for 30 min. After dehydration samples were platinum coated by the auto-fine coater (JEC-3000FC) at 20 mA current in a vacuum for 30 s. Images were acquired using JSM IT 100 JEOL (Japan) Electron Microscope using the following parameters. The tungsten is an electron beam source, with high vacuum conditions for high-resolution and visualization purposes. The images were obtained at 5 to 20 KV electron volts (depending on the sample) at a working distance of 5–10 mm from the pole piece. Modes of imaging were acquired using Secondary electron detector (SEI) images (Ahmed et al., 2022; Mirani, Fatima, et al., 2018).
Atomic Force Microscopy
Atomic force microscopy (AFM) was conducted using an Agilent − 5500 AFM system (USA) according to the following procedure. Bacterial cells were cultured on sterile silicon chips as previously described and washed to remove planktonic cells. The biofilms were then heat-fixed for 20 minutes. AFM images were captured under the ACFAM mode using a force constant of 42 N/m and a resonance frequency of 330 kHz. To study cell shape, the biofilm positive slides were first removed from the culture medium and washed twice with PBS. Vortexing at 200 rpm for 30 seconds in PBS was performed to remove the upper layers of the biofilm consortia and expose the cells directly attached to the glass surface. The dispersed cells in the buffer were then collected through centrifugation at 10,000 RPM for 5 minutes and fixed on a glass slide for further analysis. The slides were mounted on the AFM using double tape and metal discs. All morphological analysis was carried out in PBS at 37°C. For the measurements, probes were positioned under optical control at the center of the cells, and force-distance curves were acquired with a constant approach velocity of 1 µm/s.
Image Processing:
All images were processed using Gwyddion 2.60 software. The pixel dimensions of each image were set at 1280 pixels along the horizontal axis and 960 pixels along the vertical axis. The analysis was conducted using the unit channel G. The X and Y dimensions were adjusted according to the provided scale for each image. Throughout the analysis, the dimensions along the Z-axis for each sample unit were uniformly set at 1 µm. To preprocess the images, the surface slope was flattened, and data was leveled by mean plane subtraction. This was achieved by selecting the "leveling data by mean plane subtraction" command in the Gwyddion interface. The command followed the equation Z (X, Y) = a + bx + cx2 + dx3 + By + Cy2 + Dy3, which employs the least squares method to approximate surfaces with a plane. Next, data leveling was performed to ensure facet points were oriented upward. The "facet point upward" command was used to accomplish this. Additionally, individual cell images were made three points flat by applying the "three-point leveling" function. In some images, spikes were filtered using median filtration, and striated lines were removed by line-by-line leveling using the "align rows" command in the data processing interface. Any common artifacts introduced by line-by-line leveling were addressed using the masking command available in the Gwyddion tools. The "histograms adjust" command was applied to highlight the prominent features of the images. For quantitative physical characterization of surface features, the line profile command in the tools interface was utilized. This command allowed precise measurements of particle height, width, and length. To extract volume data, the volume menu toolbox in GWYDDION 2.60 offered specialized functions to generate curves or spectra in each pixel. When a stack of images along the Z-axis was not available, the software interpreted the data as a set of curves attached to each pixel in the XY plane, serving as an alternate representation of the Z-axis.