Hi Sarah,

to validate your KO, I would validate the editing (insertion) by sequencing of the genomic region using standard PCR and Sanger sequencing. You should be able to detect your insertion and show your frame-shift via an alignment. If you insertion is very early in the transcript, it should get rid of all alternative transcripts. 

If you worry about other isoforms, you could do RT-QPCR on the exon specific junction of every transcripts. If there are all absent vs. WT you were successful in KO the gene and all the alternative transcripts. 

Afterwards, you can validate that the protein is KO by doing a western blot and comparing the expression with WT cells. You should see a band in the WT and no expression in the KO cells. If you have a problem with the antibody, is there any surrogate proteins that you can look at? Have you considered a quick LC-MS/MS experiment of whole cell lysate or of a CO-IP? 

Finally, if your protein has a specific function (like a receptor, or else), I would show that KO cells are not responding the their specific stimuli. As an example, in our lab, we have KO the IFNAR1 in 293T cells. When we treat the cells with IFN-alpha, the KO cells are not stimulated by the cytokine in comparison with the WT cells. 

Altogether, these experiments will give you some genomic/proteomic and fonctional evidence that your genome editing was successful. 

Best,

Nicolas

what do you mean by gene knockout? Are cells after insertion still express truncated version of the gene which renders it non functional. In that case it is going to be difficult to assess by western. unless you have a particular antibody for the truncated region to prove your insertion worked.

or if the truncated part as enzymatic activity you need to show its nonfunctionality by enzyme assay. 

or if truncated region has specific protein binding property you do IP and show decrease binding of that protein.

good luck

Hi Sarah,

to validate your KO, I would validate the editing (insertion) by sequencing of the genomic region using standard PCR and Sanger sequencing. You should be able to detect your insertion and show your frame-shift via an alignment. If you insertion is very early in the transcript, it should get rid of all alternative transcripts. 

If you worry about other isoforms, you could do RT-QPCR on the exon specific junction of every transcripts. If there are all absent vs. WT you were successful in KO the gene and all the alternative transcripts. 

Afterwards, you can validate that the protein is KO by doing a western blot and comparing the expression with WT cells. You should see a band in the WT and no expression in the KO cells. If you have a problem with the antibody, is there any surrogate proteins that you can look at? Have you considered a quick LC-MS/MS experiment of whole cell lysate or of a CO-IP? 

Finally, if your protein has a specific function (like a receptor, or else), I would show that KO cells are not responding the their specific stimuli. As an example, in our lab, we have KO the IFNAR1 in 293T cells. When we treat the cells with IFN-alpha, the KO cells are not stimulated by the cytokine in comparison with the WT cells. 

Altogether, these experiments will give you some genomic/proteomic and fonctional evidence that your genome editing was successful. 

Best,

Nicolas

Hi Sarah

Thanks for valuable comments provided by Deenadayalan Bakthavatsalam and Nicolas Tremblay.

During the very first beginning of your experiment designing you should know almost everything about your gene like structure, function, activity etc.

Regarding your question you didn't tell anything about the insertion location [1bp]. You should know where this mutation is and how far away from the ATG, if it's nearby ATG then that gene may frame-shifted and it will not work anymore, but if its far away from ATG then that protein might be translated but it still may not work or part of it will works, so I would recommend to know everything about your gene from structure to function.

All the best.