Dear all,

I'm currently performing southern blotting to verify my CRISPR/Cas9 mediated knock-in of a 2.4 kb insert into chromosome 6 of near haploid HAP1 cells.

For that, I've isolated and purified high molecular weight DNA with a protocol, which should avoid unwanted shearing of my DNA.

For the digest, I decided to cut with EcoRI, which should give me a 6.4 kb fragment, if the insert was integrated at the desired locus and an approx. 4 kb fragment in my wild-type control cells.

The blotting process, hybridization (Church buffer) and radioactive labeling of probes, (internal and genomic) went well.

After washing and exposure, I expect single bands on the internal control blot, if the reporter is integrated only at the desired locus. Further, one band with a size of 6.4 kb (corresponding to correct, homozygous targeting events) should appear in my CRISPR/Cas9 modified cell lines but not in the wild-type control. Since we already knew that the targeting turned heterozygous, we expected 4 kb bands also in the targeted cells. However, after developing of the blots, we also saw the 6.4 kb band for targeted clones in the wild-type control but no band in the internal control.

As of now, we do not have a clue, why we are able to detect this band in the wild-type control. The genomic probe is specific to the chosen locus (I've blasted the probe afterwards to ensure that) and was purified before the radioactive labeling process. Contamination with targeted cells cannot be an issue as well, since no band on the internal control blot for the wild type cells became detectable.

Do you have any ideas, why we are able to see those band patterns in wild-type control?

Looking forward to getting some helpful answers!

I'll attach the results of the blotting as images.