DNA transposition is an important biological phenomenon that mediates genome rearrangements, inheritance of antibiotic resistance determinants, and integration of retroviral DNA. Transposition has also become a powerful tool in genetic analysis, with applications in creating insertional knockout mutations, generating gene-operon fusions to reporter functions, providing physical or genetic landmarks for the cloning of adjacent DNAs, and locating primer binding sites for DNA sequence analysis. DNA transposition studies to date usually have involved strictly in vivo approaches, in which the transposon of choice and the gene encoding the transposase responsible for catalyzing the transposition have to be introduced into the cell to be studied (microbial systems and applications are reviewed in ref. 1). However, all in vivo systems have a number of technical limitations. For instance, the transposase must be expressed in the target host, the transposon must be introduced into the host on a suicide vector, and the transposase usually is expressed in subsequent generations, resulting in potential genetic instability. A number of in vitro transposition systems (for Tn5, Tn7, Mu, Himar1, and Ty1) have been described, which bypass many limitations of in vivo systems. For this purpose, we have developed a technique for transposition that involves the formation in vitro of released Tn5 transposition complexes (TransposomesTM) followed by introduction of the complexes into the target cell of choice by electroporation. In this report, we show that this simple, robust technology can generate high-efficiency transposition in all tested bacterial species (Escherichia coli, Salmonella typhimurium, and Proteus vulgaris) We also isolated transposition events in the yeast Saccharomyces cerevisiae.

Lentivector gene therapy for X-linked chronic granulomatous disease (X-CGD) has proven to be a viable approach, but random vector integration and subnormal protein production from exogenous promoters in transduced cells remain concerning for long-term safety and efficacy. A previous genome editing-based approach using SpCas9 and an oligodeoxynucleotide donor to repair genetic mutations demonstrated the capability to restore physiological protein expression, but lacked sufficient efficiency in quiescent CD34+ hematopoietic cells for clinical translation. Here, we show transient inhibition of p53-binding protein 1 (53BP1) significantly increased (2.3-fold) long-term homology directed repair (HDR) to achieve highly efficient (80% gp91phox+ cells compared to healthy donor control) long-term correction of X-CGD CD34+ cells.

X-linked chronic granulomatous disease is an immunodeficiency characterized by defective production of microbicidal reactive oxygen species (ROS) by phagocytes. Causative mutations occur throughout the 13 exons and splice sites of the CYBB gene, resulting in loss of gp91 phox protein. Here we report gene correction by homology-directed repair in patient hematopoietic stem/progenitor cells (HSPCs) using CRISPR/Cas9 for targeted insertion of CYBB exon 1–13 or 2–13 cDNAs from adeno-associated virus donors at endogenous CYBB exon 1 or exon 2 sites. Targeted insertion of exon 1–13 cDNA did not restore physiologic gp91 phox levels, consistent with a requirement for intron 1 in CYBB expression. However, insertion of exon 2–13 cDNA fully restored gp91 phox and ROS production upon phagocyte differentiation. Addition of a woodchuck hepatitis virus post-transcriptional regulatory element did not further enhance gp91 phox expression in exon 2–13 corrected cells, indicating that retention of intron 1 was sufficient for optimal CYBB expression. Targeted correction was increased ~1.5-fold using i53 mRNA to transiently inhibit non-homologous end joining. Following engraftment in NSG mice, corrected HSPCs generated phagocytes with restored gp91 phox and ROS production. Our findings demonstrate the utility of tailoring donor design and targeting strategies to retain regulatory elements needed for optimal expression of the target gene.

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