Molecular Additives Significantly Enhance Glycopolymer-Mediated Transfection of Large Plasmids and Functional CRISPR-Cas9 Transcription Activation Ex Vivo in Primary Human Fibroblasts and Induced Pluripotent Stem Cells.

Abstract

Fast, efficient, and inexpensive methods for delivering functional nucleic acids to primary human cell types are needed to advance regenerative medicine and cell therapies. Plasmid-based gene editing (such as with CRISPR-Cas9) can require the delivery of plasmids that are large (∼9.5-13 kbp) in comparison to common reporter plasmids (∼5-8 kbp). To develop more efficient plasmid delivery vehicles, we investigated the effect of plasmid size on the transfection of primary human dermal fibroblasts (HDFs) and induced pluripotent stem cells (iPSCs) using a heparin-treated trehalose-containing polycation (Tr4-heparin). Transfections with 4.7 kbp to 10 kbp plasmids exhibited high rates of polyplex internalization with both plasmid sizes. However, transfection with the large plasmid was nearly eliminated in HDFs and significantly reduced in iPSCs. Molecular additives were used to probe intracellular barriers to transfection. Chloroquine treatments were used to destabilize endosomes, and dexamethasone and thymidine were used to destabilize the nuclear envelope. Destabilizing the nuclear envelope resulted in significantly increased large-plasmid-transfection, indicating that nuclear localization may be more difficult for large plasmids. To demonstrate the potential clinical utility of this formulation, HDFs and iPSCs were treated with to dexamethasone-Tr4-heparin polyplexes encoding dCas9-VP64, synthetic transcription activator, targeted to collagen type VII. These transfections enhanced collagen expression in HDFs and iPSCs by 5- and 20-fold, respectively, compared to an untransfected control and were the more effective than the Lipofectamine 2000 control. Functional plasmid transfection efficiency can be significantly improved by nuclear destabilization, which could lead to improved development of nonviral vehicles for ex vivo CRISPR-Cas9 gene editing.