Hello!
I am planning on using siRNA knockdown in neurones to knock down expression of ion channel X.
Alongside measurements of ion channel X function following control and targeted siRNA transfection, I would like to measure the efficiency of the knockdown, ideally using both RT-PCR and Western blotting.
I can do single-cell RT-PCR to look at mRNA levels, but how many cells would people recommend as necessary for a Western blot of ion channel X protein expression level?
siRNA will be Cy3 tagged and so I will know which cells should experience knockdown, again easy for single cell RT-PCR, but I don't really fancy culturing neurones, transfecting and then later trypsinising and subjecting them to FACS.
Thanks in advance!
Ewan
It sounds like you might want to just do some immunofuorescence-based cell imaging instead. Do you have an antibody that works for immunostaining applications?
Hi Mark, thanks, ICC would be an option I guess, but I'd be more convinced by seeing bands on gels from control vs. targeted siRNA than pixel intensities, although if the knock down was very efficient then it could be quite compelling...
Yes, especially if you see the Cy3 positive cells have less signal than Cy3 negative cells.
That's a good point, the Cy3 labelling would act as a nice internal control...thanks!
Hi Ewan,
I did siRNA knockdown of Hsc70 (yes, way more abundant) and used Western Blot together with an Odyssey Reader for quantification. The advantage is that the Odyssey measures in a linear range and does not reach saturation (or will at least tell you if it does) like your HRP signal would. You can then quantify the band intensity and get some really nice % knockdown numbers in addition to still showing the actual bands. you just need different secondary antibodies. Worked great for me.
Good luck, Susanne
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