I want to measure the effect of siRNAs on cell proliferation in a 7-day time course with a WST-1 assay.
I've done time courses before where I've applied drugs or ligands to the cells. What I normally do is plate out the cells in DMEM with 10% FCS, 1% L-glut and 1% pen/strep. I leave them to adhere overnight, and then the next day change the media to a reduced serum version (0.1%FCS) and serum-starve them for 24hours before I add the drug/ligand. Then I leave them for the 7 days before reading the proliferation.
:S I'm not sure how to apply this to my transfection with siRNA.
In a normal transfection, I would plate the cells in DMEM with 10%FCS, 1% L-glut and NO antibiotics, then refresh this media just before transfection and transfect in that, and assay for knockdown after 24 hours.
My questions are:
1. How can I apply the serum-free media to my transfection? Can I transfect in serum-free media (also lacking pen/strep), or...
2. ... do I transfect as normal and then change the media to serum-free after a period of time? (Either six hours or 24 hours after transfection).
3. Also, if I go for option 2 and change the media after transfection, should I keep the pen/strep out, or is it OK to add antibiotics AFTER the transfection has taken place?
Thanks for any help anyone can give. If it helps, I'm using Lipofectamine 2000 for transfection.
Thanks!













