Variable alterations in frequencies of CRISPR/Cas9-induced nucleotide deletion and insertion in transgenic maize callus tissues exposed to chromatin modifiers.

Given the growing emphasis on the efficient application of CRISPR/Cas9 in plant genome editing, there is a need for methodologies that enhance targeted mutagenesis efficiency. In the present study, we utilized stable transgenic maize callus tissues harboring active CRISPR/Cas9 vector systems targeting either the abnormal cytokinin response 1 (ARE1) gene or the potassium channel gene (KAT2). The presented DNA sequencing data indicate that the KAT2-targeting vector, which utilizes a 3'-end truncated guide RNA, retained mutagenic activity, albeit at a notably low frequency. We quantified mutagenic events following exposure to elevated temperatures (37 °C for 48 h) and various chromatin-modifying compounds, including sodium butyrate (NaBt), nicotinamide (NA), and 5-azacytidine (AZA). High-temperature treatment alone increased the frequency of nucleotide deletions (5.8-fold relative to the untreated samples) in the ARE1 target gene. Conversely, in the KAT2 gene, both deletions and insertions were stimulated (6.4 and 6.2-fold, respectively). As a novel approach, we evaluated the mutagenic efficacy of chromatin-modifying compounds administered in conjunction with heat treatment. The combined application of NaBt and AZA with elevated temperature was particularly effective in increasing insertion frequencies; in the ARE1 (with NaBt + heat: 17.3-fold) and KAT2 (with AZA + heat: 21.4-fold) genes. Cell cycle analysis of suspension-cultured cells revealed that chromatin-modifying treatments altered cell cycle progression, evidenced by an accumulation of cells in the G2 phase and a significant reduction in the number of mitotic cells. Subsequent DNA sequencing demonstrated a differential increase in the induction of deletions and insertions, alongside an expansion of the deleted regions.