Hi,
I recently made a mutant cell line by excising an exon from the gene, DNMT1, using CRISPR-CAS9. I isolated a single cell population that has this mutation and confirmed the mutation using PCR and Sanger sequencing. My PI also wants to use RT-qPCR to show that the sequence is missing in the mRNA. I made 3 sets of primers targetting the exon, so I would only expect amplification in the negative control cell line and not in the mutant line. However, when I ran the qPCR, I got normal amplification of this DNMT1 exon in both the negative control and the mutant line (~ct values around 23 for both).
I've extracted RNA three separate times to make sure I didnt have RNA contamination the times prior, but I still get the same result.
If anyone has experience with this or may have solutions, any help is appreciated!
Hi, Deepak,
Real-time PCR assays are sometimes tricky.
Can you please specify whether you use a Taq-man or SYBRgreen chemistry.
If you use SYBRGreen as a reporter dye, running a non-template control during your qRT-PCR experiment can be vital in order to account for any possible side contamination or primer dimers.
May I suggest you also run a classic end-point RT-PCR (not qPCR) with the primers you initially used to validate the deletion?
Maybe design your RT-PCR primers to produce a larger product in wild-type and a smaller product in the exon knockout by using primers from flanking exons. When you try to amplify a sequence that is not present, the primers will likely find something to amplify (pseudogenes, contaminating DNA etc). Of course, you can sanger sequence the products to see what you have amplified.
Did you use dual gRNA to excise the exon 1 or created in-dels with only one gRNA targeting exon 1?
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