Abstract
Nicotiana benthamiana serves as a unique platform for biopharmaceutical production, offering advantages such as efficient and scalable protein synthesis. In addition, custom N-glycans can be engineered on biopharmaceutical glycoproteins. Yet, plant-native glycosyltransferases and glycoside hydrolases need to be removed to prevent undesired modifications of tailored N-glycans. CRISPR-based systems offer tremendous potential; however, the ploidy of the allotetraploid N. benthamiana can make genome editing challenging when attempting to knock out multiple undesired enzymes using transgenes. Here, we report a highly efficient CRISPR ribonucleoprotein (RNP)-protoplast genome editing strategy for rapid, single-generation platform engineering. We delineate the editing characteristics of ErCas12a RNPs and apply hydrogel protoplast immobilization to characterize true single-cell regeneration. We target three β-hexosaminidases responsible for removing terminal GlcNAc and/or GalNAc residues from N-glycans and verify their inactivity via MALDI-TOF-MS N-glycan analysis. We achieve up to 89.6%, 95.3% and 86.5% on-target editing in the absence of off-target editing. We demonstrate the feasibility of low cell density (10(4) ml(-1)) regeneration of individual CRISPR-edited protoplasts in 12-14 weeks, carrying intended tetra-allelic and/or deca-allelic mutations while maintaining monoclonality. Despite the occurrence of genome duplications during the single-cell regeneration of N. benthamiana protoplasts, high-efficiency genome editing paired with shoot induction frequencies exceeding 89% facilitated the ubiquitous identification of desired β-hexosaminidase mutants. We anticipate that this genome-editing method will rapidly advance glyco-engineering in polyploids such as N. benthamiana.