Dear all,
I've a mouse epithelial cell line that is constituvely expressing Cas9, and I infected it with a vector that expresses Dox-inducible gRNA. Cells were selected for Cas9 expression, and 95-99% cells are infected with the inducible gRNAs as validated by flow cytometry by the use of a reporter protein.
I generated 5 different cell lines that are constitutively expressing Cas9 and I infected them with: 1)scrambled control, 2) gRNA1 against exon1, 3) gRNA2 against exon1, 4) gRNA1 against exon2, 5) gRNA2 against exon2. This 4 gRNAs are targeting the same mouse gene.
I have not found any data on the literature about the Dox concentration for that cell line. That's why I treated it for 4 days with two different doses of Dox (Doxycycline hyclate, from Sigma, reconstituted following the protocol in RNAse and sterile H20): 1 ug/ml (as reported in the paper where the plasmids were described) and 10 ug/ml. I changed the media every other day. My cells looked fine: I did not get more cells floating in the media and they did not change the morphology.
Upon 4 days treatment with Doxycycline, I isolated DNA and I followed the instructions of the Genome Editing Detection Kit from IDT to detect missmatches. The internal homoduplex and heteroduplex controls of the kit are working. Unfortunately I do not see heteroduplexes formation in any of the above-mentioned conditions for 4 gRNAs designed targeting the same gene. I'm not sure if it is due to the Dox concentrations, to the time treatment, or just because my gRNAs are not working...
Regarding the last possibility, it is me who designed the gRNAs by using the crispr.mit.edu webpage from Zhang lab. I introduced the coding sequence of two different exons of interest, and then I chose 2 gRNAs per exon that were in the coding strand. It is also me who designed the primers for the T7E1 endonuclease assay, and I validated them by genomic DNA amplification (to check that only one PCR product is produced). That said, they should work. Obviously some of them may not work but I think it would be quite unlikely that the 4 would not work... What's your point on that?
I would treat the different cell lines with different Dox doses such as 0,1 ug/ml, 10 ug/ml, 25 ug/ml, 100 ug/ml, for 5 days and I will change the media daily. Then I will isolate the DNA and hopefully then I will detect some missmatches. Does somebody have a better idea about how I could have it working? Could it be that this cell line is non-responding to doxycycline at all?
I would be really grateful if somebody could give me some kind of help :) :)
Thank you,
Anna
Hi Anna,
First, you should check whether your sgRNAs are indeed expressed upon doxycyline treatment. This can be done by small RNA northern blotting or qRT-PCR.
Second, you can check the efficacy of your sgRNAs by in vitro cleavage assay or by transfecting them into easy-to-transfect cell lines such as HEK293 (of course you should co-transfect Cas9 in this case).
Cheers,
I also facing the same condition with you and did u solve the problems??
and can anyone give me advise .... thanks Lewis....
Dear Dooyoung and Kyaw,
Thanks for your input. Indeed I isolated the DNA and performed with it an in vitro cleavage assay in these cancer cell lines, and I only saw the WT band in all my different conditions.
I will repeat this procedure in case something went wrong.
I will keep you posted.
Best,
Anna
Hi Anna!
it might be worth having a look at this paper: Article An iCRISPR Platform for Rapid, Multiplexable, and Inducible ...
"iCas9 hPSCs were treated with doxycycline (2 μg/ml) for one or two days before and during transfection. "
we have used this method, and it works really well to induce Cas9 expression upon addition of doxycycline. You'll probably need to optimise this for your cell lines, but this could be a good starting point.
Good luck!
Hi Dani :)
Thank you a lot. It seems to be an interesting paper, I will give a closer look.
Good luck for you too, I hope everything is going well!
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