How much blast or Dox did you use? (3ul isn’t particularly helpful, what was your final concentration).

Remember Dox and Blast are both antibiotics so at the end of the day will kill cells if the concentration is high enough. I’d suggest performing a kill curve. There are several ways to do this but the easiest ‘cheat’ way to do it would be to seed your cells into 24 well plates and add varying amounts of Dox and Blast separately. Look to see which cells have the highest concentration of Blast and Dox without being killed/messed up in the process and use those concentrations in your next experiment.

Also I assume these cells have a Tet- inducable plasmid containing Blast resistance? Assuming your induced protein isn’t causing cell death, you could perform a kill curve on the non-transduced cells and use the lowest concentration of blasticidine that causes 100% cell death.

Finally, is the blast resistance under a separate promoter to your inducible protein? Usually the blast resistance is under a constant promoter so you select your cells using blast (which is a bit of a shock) then after the appropriate amount of time needed to kill the non transduced cells, you add dox.

All the Best!

Hi Jack,

The concentration for Doxy and blasti were 1 mg/mL and 10 mg/mL respectively. These concentrations have been standardized in the lab and has been used for very long time. But, this time we observed all the cell dying

Okay, did you treat a colony from a single clone or did you treat the whole population? If you’ve tested out those concentrations before then the only thing I can think of at the moment is that the transduction efficiency (for what ever reason) was much lower than you expected.

I would suggest maybe treating some cells with Dox and not Blast in order to determine the %of cells that took up the plasmid. Can you perform some kind of fluorescence/FACS? If only 10% integrated then it makes sense that Blast would kill the 90%

(This is assuming you treated a population, not a clone)