CRISPR/Cas9-mediated HDR-induced Knock-in (KI) in zebrafish seems to yield very low efficiency in germ-line transmission. To obtain a higher efficiency, we may need to optimize several factors such as the type of the template, homology arm lengths, etc. Thus, I am planing to compare different types of templates (ssODN, dsDNA, Plasmid) seeking which template would give a higher KI efficiency quantitatively. I would microinject donor DNA, Cas9 protein and gRNA together, and after 24 hpf, check the KI events by PCR. However, I was wandering if I can evaluate KI events quantitatively either by RT-qPCR or by another method. Has anyone tried such experiment or has any comments on this? I really appreciate your comments and suggestions.
I am doing KI in plant protoplasts using ssDNA as donor templates for HDR and evaluating them via Sanger Sequencing after multiplication.
I was using Sanger sequencing in combination with the ICE analysis tool (https://www.synthego.com/products/bioinformatics/crispr-analysis). It gives you an estimate of editing events. However, it is not very reliable for knock-ins.
You could try to introduce a unique restriction site by generating a synonymous change in the the template. Thereby, you could run a PCR, followed by restriction digest.
I normally introduce the restriction enzyme site into the DNA donor (ssDNA, dsDNA, or so on). After editing, you can do PCR and digest your PCR product with restriction enzyme. Band quantification should be fine for estimate the KI efficiency.
And, of course, you could also use TIDE, TIDER and Inference of CRISPR Edits (ICE) to test it.
Thank you for your responses. Ngoc Tung Tran in my case, KI allele copy number is very minor and so in PCR with primers flanking the KI site does not amplify the KI allele rather the WT allele. My insert is ~200 bp (without homology arms), so it should be very easy to distinguish if amplified. However, I can see bands if I use a primer specific to the insert. That is why I thought to try q-PCR. Hope I can optimize a strategy.