I've been doing some experiments with my siRNAs, and have had good success in observing knock down by Western Blotiing after 24 hrs of transfection: my test siRNAs give good knockdown, while mock/untransfected and scrambled controls have no observable effect.
But now that I'm applying them in cell proliferation assays, I'm having problems. I can still get good knockdown with my test siRNAs, but the scrambled ones are also killing/inhibiting my cells! Even more than the test siRNAs in some cases
Does anybody have any advice/experienced this before?
Can scrambled siRNAs have general non-specific toxic effect? I hope not, because I can't really afford to buy any more!
Some background:
-- I'm transfecting with Lipofectamine 2000.
-- I've done a range of siRNA concentrations (8, 16, 24 and 32nM), keeping the siRNA:lipofectamine ratio the same throughout. I'm not observing much cell death/inhibition in the mocks, so I don't think the amount of lipofectamine is the problem?
-- Cells are prepared by seeding in pen-strep free DMEM with 10%FCS, adhereing overnight, transfecting the next day, changing media after 6 hrs (to DMEM with 0.1% FCS and no pen-strep) and leaving for 7 days to proliferate before reading.
Thanks for any help anybody can give. I know I'm on this board a lot moaning about my silly siRNAs!
Edited by steffi333, 04 May 2011 - 05:29 AM.














