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Abstract
Genome-editing is being implemented in increasing number of plant species using engineered sequence specific nucleases (SSNs) such as Clustered Regularly Interspaced Short Palindromic Repeats/CRISPR-associated systems (CRISPR/Cas9), Transcription activator like effector nucleases (TALENs), and more recently CRISPR/Cas12a. As the tissue culture and regeneration procedures to generate gene-edited events are time consuming, large-scale screening methodologies that rapidly facilitate validation of genome-editing reagents are critical. Plant protoplast cells provide a rapid platform to validate genome-editing reagents. Protoplast transfection with plasmids expressing genome-editing reagents represents an efficient and cost-effective method to screen for in vivo activity of genome-editing constructs and resulting targeted mutagenesis. In this study, we compared three existing methods for detection of editing activity, the T7 endonuclease I assay (T7EI), PCR/restriction enzyme (PCR/RE) digestion, and amplicon-sequencing, with an alternative method which involves tagging a double-stranded oligodeoxynucleotide (dsODN) into the SSN-induced double stranded break and detection of on-target activity of gene-editing reagents by PCR and agarose gel electrophoresis. To validate these methods, multiple reagents including TALENs, CRISPR/Cas9 and Cas9 variants, eCas9(1.1) (enhanced specificity) and Cas9-HF1 (high-fidelity1) were engineered for targeted mutagenesis of Acetolactate synthase1 (ALS1), 5-Enolpyruvylshikimate- 3-phosphate synthase1 (EPSPS1) and their paralogs in potato. While all methods detected editing activity, the PCR detection of dsODN integration provided the most straightforward and easiest method to assess on-target activity of the SSN as well as a method for initial qualitative evaluation of the functionality of genome-editing constructs. Quantitative data on mutagenesis frequencies obtained by amplicon-sequencing of ALS1 revealed that the mutagenesis frequency of CRISPR/Cas9 reagents is better than TALENs. Context-based choice of method for evaluation of gene-editing reagents in protoplast systems, along with advantages and limitations associated with each method, are discussed.
Highlights
Genome-editing by engineered sequence specific nucleases (SSNs) is a technological breakthrough that enables precise alterations to DNA, representing a new frontier in genetics
Single guide RNAs and Transcription activator like effector nucleases (TALENs) were designed for targeted mutagenesis of Acetolactate synthase1 (ALS1) (Butler et al, 2015), Enolpyruvylshikimate- 3-phosphate synthase1 (EPSPS1) (Figures 1A,B) and their paralogs, ALS2 and EPSPS2 respectively, in potato (Supplementary Table S3)
For detection of successful delivery and expression of the genome-editing reagents, green fluorescent protein (GFP) was co-delivered on the same plasmid as the Cas9 and TALEN expression cassettes (Figure 1C)
Summary
Genome-editing by engineered sequence specific nucleases (SSNs) is a technological breakthrough that enables precise alterations to DNA, representing a new frontier in genetics. EvoCas (evolved Cas9) (Casini et al, 2018) have been designed based on structure-guided protein engineering to reduce non-specific DNA interactions thereby minimizing genome-wide off-targets. Genome-editing reagents are delivered into plant cells via Agrobacterium-mediated transformation, protoplast transfection, or particle bombardment and typically, selection is employed to regenerate plants with integrated constructs, which intend to induce desired mutations (Yin et al., 2017). Plant transformation and regeneration processes from engineered cells and tissues typically require long and tedious tissue culture procedures. Protoplast transformation is a direct means to deliver genome-editing reagents and does not require a biological vector. Protoplasts can be used to detect reporter genes by microscopy and are amenable to cell sorting (Xing and Wang, 2015)
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