Present adoptive immunotherapy strategies are based on the re-targeting of autologous T-cells to recognize tumor antigens. As T-cell properties may vary significantly between patients, this approach can result in significant variability in cell potency that may affect therapeutic outcome. More consistent results could be achieved by generating allogeneic cells from healthy donors. An impediment to such an approach is the endogenous T-cell receptors present on T-cells, which have the potential to direct dangerous off-tumor antihost reactivity. To address these limitations, we assessed the ability of three different TCR-α-targeted nucleases to disrupt T-cell receptor expression in primary human T-cells. We optimized the conditions for the delivery of each reagent and assessed off-target cleavage. The megaTAL and CRISPR/Cas9 reagents exhibited the highest disruption efficiency combined with low levels of toxicity and off-target cleavage, and we used them for a translatable manufacturing process to produce safe cellular substrates for next-generation immunotherapies.

Chimeric antigen receptor (CAR) T cell therapy represents the first U.S. Food and Drug Administration approved gene therapy and these engineered cells function with unprecedented efficacy in the treatment of refractory CD19 positive hematologic malignancies. CAR translation to solid tumors is also being actively investigated; however, efficacy to date has been variable due to tumor-evolved mechanisms that inhibit local immune cell activity. To bolster the potency of CAR-T cells, modulation of the immunosuppressive tumor microenvironment with immune-checkpoint blockade is a promising strategy. The impact of this approach on hematological malignancies is in its infancy, and in this review we discuss CAR-T cells and their synergy with immune-checkpoint blockade.

Programmable gene integration in human cells has the potential to enable mutation-agnostic treatments for loss-of-function genetic diseases and facilitate many applications in the life sciences. CRISPR-associated transposases (CASTs) catalyze RNA-guided DNA integration but thus far demonstrate minimal activity in human cells. Using phage-assisted continuous evolution (PACE), we generated CAST variants with >200-fold average improved integration activity. The evolved CAST system (evoCAST) achieves ~10 to 30% integration efficiencies of kilobase-size DNA cargoes in human cells across 14 tested genomic target sites, including safe harbor loci, sites used for immunotherapy, and genes implicated in loss-of-function diseases, with undetected indels and low levels of off-target integration. Collectively, our findings establish a platform for the laboratory evolution of CASTs and advance a versatile system for programmable gene integration in living systems.

Mucopolysaccharidosis type I (MPS I) is a lysosomal storage disease characterized by mutations to the α-L-iduronidase (IDUA) gene resulting in inactivation of the IDUA enzyme. The loss of IDUA protein results in the progressive accumulation of glycosaminoglycans within the lysosomes resulting in severe, multi-organ system pathology. Gene replacement strategies have relied on the use of viral or nonviral gene delivery systems. Drawbacks to these include laborious production procedures, poor efficacy due to plasmid-borne gene silencing, and the risk of insertional mutagenesis. This report demonstrates the efficacy of a nonintegrating, minicircle (MC) DNA vector that is resistant to epigenetic gene silencing in vivo. To achieve sustained expression of the immunogenic IDUA protein we investigated the use of a tissue-specific promoter in conjunction with microRNA target sequences. The inclusion of microRNA target sequences resulted in a slight improvement in long-term expression compared to their absence. However, immune modulation by costimulatory blockade was required and permitted for IDUA expression in MPS I mice that resulted in the biochemical correction of pathology in all of the organs analyzed. MC gene delivery combined with costimulatory pathway blockade maximizes safety, efficacy, and sustained gene expression and is a new approach in the treatment of lysosomal storage disease.

Disruption of CCR5 or CXCR4, the main human immunodeficiency virus type 1 (HIV-1) co-receptors, has been shown to protect primary human CD4⁺ T cells from HIV-1 infection. Base editing can install targeted point mutations in cellular genomes, and can thus efficiently inactivate genes by introducing stop codons or eliminating start codons without double-stranded DNA break formation. Here, we applied base editors for individual and simultaneous disruption of both co-receptors in primary human CD4⁺ T cells. Using cytosine base editors we observed premature stop codon introduction in up to 89% of sequenced CCR5 or CXCR4 alleles. Using adenine base editors we eliminated the start codon in CCR5 in up to 95% of primary human CD4⁺ T cell and up to 88% of CD34⁺ hematopoietic stem and progenitor cell target alleles. Genome-wide specificity analysis revealed low numbers of off-target mutations that were introduced by base editing, located predominantly in intergenic or intronic regions. We show that our editing strategies prevent transduction with CCR5-tropic and CXCR4-tropic viral vectors in up to 79% and 88% of human CD4⁺ T cells, respectively. The engineered T cells maintained functionality and overall our results demonstrate the effectiveness of base-editing strategies for efficient and specific ablation of HIV co-receptors in clinically relevant cell types.