The capability to apply a precise genome editing in cells remains a well-established desire. However, canonical CRISPR-Cas9 based tools introduce double-strand DNA breaks (DSBs) which can lead to spontaneous INDELs at target site [1]. Prime editing (PE) is the most recent molecular system which consists of a Cas9 nickase fused to a reverse transcriptase. PE is guided by a prime editing guide RNA (pegRNA) which codes both: a guidance to genomic target site and a matrix for the desirable edit [2]. Therefore, PE is able to mediate a targeted mutagenesis without need of DSBs or donor template.
To obtain a desirable base conversion at target site, in this work we adapted and optimized the most recent PE system for genome editing in murine embryonic stem cells. Firstly, plasmids bearing 9 distinct pegRNAs coding SatI restriction site were constructed. Next, the efficiency of genome editing was evaluated by SatI restriction analysis of PCR products which were prepared from cells transfected with plasmids coding PE and corresponding pegRNAs, respectively. Importantly, the analysis revealed that all of selected pegRNA variants showed a significant genome prime editing at the target site. To determine the status of programmable mutagenesis in separate cells, 340 single cell clones were firstly obtained by serial cell dilution. Restriction analysis of each clone unveiled that 27.9% of them obtained a target mutation. Finally, DNA sequencing analysis confirmed that all of the selected 8 clones were bearing a precisely corrected allele and no additional spontaneous mutations were observed in all analysed clones.
In summary, prime editing is a precise and efficient tool for targeted genome mutagenesis in murine embryonic stem cells.