Hi all,
I am using FuGENE to transiently transfect RPE1 cells for imaging. I usually seed them on coverslips and transfect them when they have reached 60-70% confluence. For 24-well dishes I use 30 uL OptiMEM, 500 ng DNA and 1.5 uL FuGENE. I normally fix the cells 2 days after transfection. Unfortunately, my plasmid does not express a reporter, so I cannot check for transfection efficiency before fixing.
My transfection efficiency is extremely low (
Thank you for your suggestions, Philippe!
I am trying to express a FLAG-tagged golgin consisting of about 2,000 amino acids in a pCMV vector (https://www.snapgene.com/plasmids/mammalian_expression_vectors/pCMV-Tag_2B). It unfortunately does not express any reporter.
Not sure what you mean with pure OptiMEM. I prepare a tube with the OptiMEM, add the DNA, then FuGene, let incubate 15-20 min and transfer the mix to cells in dishes that contain DMEM F-12. This general procedure works very well when I use another plasmid and HEK293T cells, so I assume the composition etc should be fine.
I'm not 100% sure about the efficiency, but I know that people have done the transfection with the exact same plasmid and GOI before (I think they mentioned the low expression, but not that it was this bad).
I tried another round of transfection and will get the results on saturday, if they come out as bad as before, I might try lowering DNA concentration or ask other labs for different transfection reagents :-). Thanks again!
Best,
Laura
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