HeLa cells (6-well) transfected with 4ug pDNA
Cells are lysed with NP40 Lysis Buffer with RNaseOUT
Protein isolated from lysates shows expression of transgene by WB
RNA isolated from lysates with Trizol and cDNA is synthesized using 1ug RNA. 260/280= 1.95 on average.
We use GoTaq mix to amplify 2uL (out of 25uL) of cDNA template.
Plasmid +ve control for transgene always works.
Transgene amplicons are barely visible after 32 cycles, control plasmid template is all blown out.
I assume because of the strong promoter, a few copy of transgene RNA is able to generate huge amount of protein.
If you are confident that the band in WB corresponds to your transgene one possible explanation is then accumulation of the protein even if the transcription if very low. If you are not sure you have to use some controls to verify the nature of the band you are detecting by WB.
Wishing you the best
Thanks for your help, I think the most likely explanation is low transfection efficiency, but I was just surprised because I always thought PCR was extremely sensitive and if you can detect your transgene by western blot then for sure PCR would be able to detect the mRNA, but perhaps Attila's point comes into play here that few transcripts can be translated many times over.
Maybe your transfections with your GAPDH construct aren't working well, but your westerns are detecting endogenous GAPDH, which is abundantly expressed by most cell lines. This scenario would explain the PCR and western results. Would your westerns differentiate between endogenous and transfected GAPDH? You might want to resequence the transfection construct to make sure that the sequence is good as well
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