CRISPR-Cas9 holds promise for treating genetic disease, but rare off-target mutations and structural variants remain as key safety concerns, especially at scales relevant to therapy. Here, we establish workflows to resolve Cas9 off-target activity in vitro at single-cell resolution and in vivo across different tissues. Using clonally expanded electroporated mouse embryos and embryonic stem cells, we reveal that individual cells exhibit unique off-target and translocation profiles, including events missed in bulk analyses. Integrating single-cell editing with chromatin accessibility, transcription, and DNA methylation measurements suggest that sequence-independent features modulate Cas9 access and cleavage, with preferential editing in regions characterized by open chromatin and lower methylation. In Cas9-inducible mouse models, editing analyses reveal organ-distinct off-target spectra, DNA repair pathway usage, indel patterns, and markedly varying translocation propensity between tissues. These findings demonstrate that off-target activity is heterogeneous across cells and context-dependent across organs, motivating sensitive single-cell analyses and organ-specific evaluation in preclinical development to more accurately assess risk and improve the safety of CRISPR-based genomic medicines.

Acute myeloid leukemia is a genetically and cellularly heterogeneous disease. We characterize 120 AMLs using genomic and transcriptomic analyses, including single-cell RNA sequencing. Our results reveal an extensive cellular heterogeneity that distorts the bulk transcriptomic profiles. Selective examination of the transcriptional signatures of >90,000 immature AML cells identifies four main clusters, thereby extending current genomic classification of AML. Notably, NPM1-mutated AML can be stratified into two clinically relevant classes, with NPM1class I associated with downregulation of MHC class II and excellent survival following hematopoietic stem cell transplantation. NPM1class II is instead associated with resistance to allogeneic T cells in an ex vivo co-culture assay, and importantly, dismal survival following hematopoietic stem cell transplantation. These findings provide insights into the cellular state space of AML, define diagnostic entities, and highlight potential therapeutic intervention points. The clinical outcomes and treatment responses of acute myeloid leukemia (AML) patients are heterogeneous. Here, the authors use bulk and single-cell sequencing approaches and identify two transcriptomic subtypes within NPM1-mutated AML with distinct immune evasion properties and responses to hematopoietic stem cell transplantation.

Mucopolysaccharidosis (MPS) IVA is a bone-affecting lysosomal storage disease (LSD) caused by impaired degradation of the glycosaminoglycans (GAGs) keratan sulfate (KS) and chondroitin 6-sulfate (C6S) due to deficient N-acetylgalactosamine-6-sulfatase (GALNS) enzyme activity. Previously, we successfully developed and validated a CRISPR/nCas9-based gene therapy (GT) to insert an expression cassette at the AAVS1 and ROSA26 loci in human MPS IVA fibroblasts and MPS IVA mice, respectively. In this study, we have extended our approach to evaluate the effectiveness of our CRISPR/nCas9-based GT in editing human CD34+ cells to mediate cross-correction of MPS IVA fibroblasts. CD34+ cells were electroporated with the CRISPR/nCas9 system, targeting the AAVS1 locus. The nCas9-mediated on-target donor template insertion, and the stemness of the CRISPR/nCas-edited CD34+ cells was evaluated. Additionally, MPS IVA fibroblasts were co-cultured with CRISPR/nCas-edited CD34+ cells to assess cross-correction. CRISPR/nCas9-based gene editing did not affect the stemness of CD34+ cells but did lead to supraphysiological levels of the GALNS enzyme. Upon co-culture, MPS IVA fibroblasts displayed a significant increase in the GALNS enzyme activity along with lysosomal mass reduction, pro-oxidant profile amelioration, mitochondrial mass recovery, and pro-apoptotic and pro-inflammatory profile improvement. These results show the potential of our CRISPR/nCas9-based GT to edit CD34+ cells to mediate cross-correction.