I would like to establish PC3 stable GFP by CRISPR KI. I’ve tried the following protocol but cells didn’t live after transfection, even they express GFP.

1. Seed cells 2*10^5/well in 6 well and incubate 24 h in DMEM containing 10%FBS and 1% P/S.

2. Prepare OPTI-MEM-lipofactamine-DNA mixture following the manufacturer instruction or papers.

a. Add 250 ul mixture directly into well and incubate for 2 or 1 day (lipofactamine2000 or 3000).

b. Prepare mixture 600 ul/well. Discard DMEM, wash with PBS twice and Add mixture incubate for 9 h (lipofactamine2000). (I found paper showed 3 h but I saw little cells with GFP so I prolong the incubation time)

3. After transfection, replace medium with DMEM containing 4 ug blasticidin. Change medium every 2 day.

After transfection, I could see GFP expressed cells but they were deattached and the rest attached cell was very little. After antibody selection for twice almost none cell was alive.

In protocol b. cells were alive but 16 h after replacing medium, transfected-well showed much more dead cells than negative control (Without plasmids and donor frag, lipofactamine only). And GFP cells were still very few and seemed to be deattached either.

The GFP was knocked in downstream of housekeeping gene CDS. I choose 3 genes and all of them showed same problem. I’ve used the same method to establish 4T1 and F10 but everything went well and I got successfully stable clone.

Is there anything I should modify in my protocol or should I give up lipofactamine and use electro transfection instead…?

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