New in vivo delivery technologies are urgently needed that enable selective genome editing of somatic cells without the limitations of existing viral delivery systems or lipid nanoparticles. We propose to develop two complementary strategies. First, by tethering Cas9 and base editor ribonucleoproteins (RNPs) to homing moieties, such as antibodies or nucleic acid aptamers, we will develop delivery systems capable of editing a specific population of target cells. As a second approach, we will engineer viral like particles (VLPs) to facilitate efficient, tissue and cell specific delivery of genome editing agents. In the process, we will develop delivery systems that are capable of targeting hematopoietic stem and progenitor cells (HSPCs), among other tissues. To evaluate the efficiency and cell-type specificity of our proposed delivery methods, we will also generate a reporter mouse that quantitatively and sensitively reports genome editing from base editors or programmable nucleases without requiring DNA sequencing. In this proposal, we intend to: (1) Design targeted ribonucleoprotein conjugates that selectively bind, enter, and edit target cells. Cell and tissue selective Cas9 and base editor RNP delivery systems will be designed by tethering genome editing proteins, directly or indirectly, to aptamer and antibody targeting moieties. The kinetics, magnitude, and specificity of RNP endocytosis, endosomal escape, and nuclear transport will be defined and genome editing efficiency and targeting specificity determined in vitro and in vivo. (2) Engineer ribonucleoprotein nanoparticle delivery systems for cell and tissue targeted genome editing. SV40 capsid proteins will be engineered to form viral like particles (VLPs) that are capable of packaging ribonucleoproteins, rather than DNA. The stoichiometry of VLP-RNP delivery systems, which affords optimal cell uptake, endosomal escape, and nuclear transport will be defined. Targeting specificity, as determined by viral capsid tropism will be defined, and genome editing efficiency analyzed in vitro and in vivo. (3) Develop a reporter mouse for facile assessment of targeted genome editing efficiency and cell- and tissue-type specificity. We will optimize a reporter gene to independently detect base editing, end-joining, or homology-directed repair. The reporter will be integrated into the Rosa26 safe harbor locus in C57BL/6 mouse embryonic stem cells to generate transgenic mice. Genome editing outcomes will be evaluated by fluorescence and luminescence measurements and correlated with high throughput DNA sequencing. (4) Demonstrate safe and effective delivery of genome editing agents in non-human primates. The delivery of genome editors to HSPCs and other target tissues will be assessed in rhesus macaques. Both mammalian and non-mammalian systems will be evaluated to optimize large scale production of the genome editor and related RNP delivery components. Targeting specificity and genome editing efficiency, as well as safety will be analyzed in vivo. We anticipate identifying effective delivery systems suitable for clinical trials.
The difficulty of delivering genome editing agents into many types of cells in animals and patients is a major challenge that must be overcome to realize their full potential to cure genetic diseases. We propose to develop two new strategies for the delivery of genome editing agents into animals and patients that will increase editing efficiency, target cell selectivity, and DNA specificity, as well as a new tool to rapidly and sensitively evaluate the delivery of these agents in mice with minimal effort and expense. These developments will advance the safety and efficacy of genome editing methods for clinical development.