Whole genome sequencing of Salmonella enterica serovar Typhimurium directed evolution experiment, involved continuous exposure of planktonic cells or biofilms to subinhibitory concentrations of acriflavine or phenylalanine-arginine β-naphthylamide (PABN)
Biofilms were grown on four 5mm glass beads (Sigma) in LB broth without salt, supplemented with 2 µg/mL acriflavine or 8 µg/mL phenylalanine-arginine beta-naphthylamide (PAβN). Untreated biofilm controls and treated planktonic controls were done in parallel. All cultures were incubated with slow shaking at 100 rpm at 30 ºC. Every 48-72 hours, one bead was removed and placed into fresh media containing 3 sterile beads. This allowed cells from the biofilm to disperse to colonies new beads and complete the full biofilm life cycle. For planktonic controls, approximately 10^5 CFU/mL was passaged into sterile treated media. After 1, 5 or 10 passages (corresponding to 2, 17 and 35 days), biofilms were removed from the three beads remaining in the culture. This was done by gently rinsing the beads in PBS to remove planktonic cells and then vortexing for 30 seconds in 1 mL PBS. This was grown overnight in 1 mL LB broth without salt, after which 250 µL was pelleted for DNA extraction and the remaining 750 µL was combined with 250 µL 40% glycerol and stored at -20 ºC until use. Planktonic cultures were prepared for DNA extraction and stored in the same way. Frozen cultures were used to inoculate sterile LB broth without salt for phenotyping assays. To get a deeper understanding of the composition of these populations, biofilm populations from passages five and ten were streaked from the frozen stock onto LB agar without salt. Three colonies were selected at random, cultured overnight in LB broth and stored and prepared for DNA extraction the same as previously described. DNA was extracted using a Quick-DNA 96 kit (Zymo) following manufacturer's instructions DNA was quantified using a Quant-iT™ high sensitivity dsDNA assay kit (Invitrogen) and normalised to 0.5 ng/µL with 10 mM Tris-HCl. From this, 2 µl of normalised DNA (1 ng total) was mixed with 3 µL Tagmentation mix, made by mixing 0.9 µL TD Tagment DNA Buffer (Illumina) with 0.09 µL TDE1 Tagment DNA Enzyme (Illumina) and 2.01 µL nuclease-free water in a chilled 96-well plate. This was then heated to 55 °C for 10 min in a PCR machine. A PCR master mix was made using 4 µL kapa2G buffer, 0.4 µL dNTPs, 0.08 µL polymerase all from the Kap2G Robust PCR kit (Merck) and 4.52 µL nuclease-free water per sample and 11 µL was added to each well of a 96-well plate. After this, 2 µL each of P7 and P5 of Nextera XT Index Kit v2 index primers (Illumina) were added to each well. Finally, the 5 µL Tagmentation mix was added to this and mixed. The PCR was run at 72 °C for 3 minutes, 95 °C for 1 minute, 14 cycles of 95 °C for 10 seconds, 55 °C for 20 seconds and 72 °C for 3 minutes. Following the PCR reaction, the libraries were quantified as previously and pooled in equal quantities. This pool underwent double-SPRI size selection between 0.5 and 0.7X bead volumes using KAPA Pure Beads (Roche). The final pool was quantified again as previously and run on a High Sensitivity D1000 ScreenTape (Agilent) using the Agilent Tapestation 4200 to calculate library pool molarity.