Which to use for standard curve? - (Mar/06/2009 )
I want to detect if a gene's expression is reactivated when I knockdown the expression of my target gene but i'm not sure what sample to use for my standard curve. Basically, I have RNA samples from cancer cells which i have transfected with an siRNA to knockdown the expression of my gene A. I have done RT-PCR and confirmed that this has been successful & i have 80-90% suppression. My hypothesis is that when gene A is knocked down, gene B will start to be expressed again (as gene A represses gene B ). But how do I confirm this by RT-PCR? I used my non-targeting siRNA samples as my standard curve to check for the suppression of gene A but I cant use these samples for the standard curve of gene B as it wont be expressed due to the fact that gene A is repressing it. I have RNA samples for 24, 48 & 72 hours post transfection all of which have 80-90% suppression of gene A. Could I use one of these for my standard curve?
I'm sorry if this is a bit confusing.
Thanks in advance for any help.
You need to use samples that you know express gene B first to determine amplification efficiency. Do you know that gene B is activated? Have you tried Westerns first to assess whether there is a change in gene B expression at the protein level when you knock down gene A? qRT-PCR may be jumping the gun. Remember, only changes at the protein level impact physiology and mRNA levels can change without changes in protein or with opposing changes in protein expression (ie: mRNA level goes up, protein goes down as is the case for proteins such as the AHR and HIF1a following activation). I'd look at Westerns first, and then if there was activation of B, then seek to quantify that change by qRT-PCR.
Dr Teeth on Mar 6 2009, 04:48 PM said:
I have the protein samples in the freezer so I'll get some Westerns running as suggested! Thanks for the advice.