I am doing flow cytometry for cell cycle with E. coli after induction of proteins that cause cell death. The procedure I followed consists in growing cells to an OD of around 0.2 take a pre induction sample, induce protein expression and samples at different time points. Cells are kept on ice and they are fixed with ethanol previously placed at -20C. Ethanol is added to a final conc of 70% dropwise with sonication in water or light vortexing and cells are let to fix (I have tried 1h, 2h or overnight). Cells are spun down and washed (I have used PBS+0.5%BSA, TBS and 50mM sodium citrate) and then treated with RNAse A in same buffer for 4h at 50C. cells are diluted to 1x10^6 CFU/mL and stained (tried several times with SYTOX green 2.5uM or Propidium iodide 10ug/mL with 30min-2h incubation at RT for SYTOX or overnight at 4C for PI) and imaged. iced water bath sonication is used to avoid clumping and light vortex is used for mixing (around 4/10). The issue is I always see pellet but most of the time the amount of events is very low even when adjusting FSC and SSC and when I am able to get signal the peaks do not show all the transitions even for the asychronous control (pleas see attached) Do we use a higher CFU count for bacteria compared to mammalian cells? Am I losing DNA during fixation? What can be improved?