Staphylococcus aureus Cas9 (SaCas9) is smaller than the widely used Streptococcus pyogenes Cas9 (SpCas9) and has been harnessed for gene therapy using an adeno-associated virus vector. However, SaCas9 requires a longer NNGRRT (where N is any nucleotide and R is A or G) protospacer adjacent motif (PAM) for target DNA recognition, thereby restricting the targeting range. Although PAM-relaxed Cas9 variants have been developed, expanded targeting is often accompanied by compromised target specificity. Here, we report the rational engineering of eSaCas9-NNG, a SaCas9 variant that recognizes relaxed NNG PAMs while maintaining high target fidelity, thereby overcoming a fundamental trade-off in Cas9-based genome editing. eSaCas9-NNG efficiently induces indels and base conversions at endogenous sites bearing NNG PAMs in human cells and mice, with editing efficiencies comparable to those of other PAM-relaxed nucleases, including SpRY, SpG, and iGeoCas9, but with reduced off-target activity. We further determine the cryo-electron microscopy structures of eSaCas9-NNG in five distinct functional states, revealing the structural basis for its relaxed PAM recognition, improved target specificity, and nuclease activation. Overall, our findings demonstrate that eSaCas9-NNG could be used as a versatile genome editing tool for in vivo gene therapy, and improve our mechanistic understanding of the diverse CRISPR-Cas9 nucleases.

CRISPR-based genome editing therapeutics are entering the clinic, offering transformative potential but also presenting potential risks. Preclinical-to-clinical toolkits are needed to assess the safety and efficacy of these new therapies and accelerate progress. Emerging technologies to monitor the biological effects of genome editors cover a range of biological scales, from the direct measurement of editing outcomes in DNA, to human microphysiological systems, and non-invasive in vivo imaging. Measurements of on-target and off-target editing outcomes, including sequences unique to humans, provide essential benchmarks to understand functional responses. Microphysiological systems, including organoids and organs-on-chips, enable phenotypic evaluations of editing strategies in varied organ lineages and disease states. Non-invasive imaging modalities can track the biodistribution and activities of genome editors and edited cells in vivo. Collectively, these technologies provide complementary insights across different scales, from the single nucleotide to the whole organism, bridging preclinical therapeutics development with clinical trials. CRISPR-based genome editing therapeutics are entering the clinic, but in vitro and in vivo tools are needed to assess their safety and efficacy. The authors review complementary technologies to monitor the biological effects of genome editing across scales, including the direct measurement of editing outcomes in DNA, human microphysiological systems and non-invasive in vivo imaging.

The microenvironment in solid tumors represents an immunosuppressive therapeutic barrier to CAR T cell therapy, and it is currently unknown whether it can be reshaped by the deletion of negative regulators in CAR T cells. To address this knowledge gap, we evaluated the intrinsic and extrinsic effects of deleting the negative regulator Regnase-1 (Reg-1) in B7-H3-CAR T cells for the immunotherapy of osteosarcoma. Reg-1 knockout (KO) improved the antitumor activity of human and murine B7-H3-CAR T cells in vivo. In immune-competent models, Reg-1 KO also endowed murine B7-H3-CAR T cells with the ability to create a proinflammatory landscape characterized by an influx of interferon gamma (IFN-γ)-producing endogenous T cells and natural killer (NK) cells and a reduction of inhibitory myeloid cells, including M2-like macrophages. Thus, deleting Reg-1 has cell- and non-cell-autonomous benefits, nominating Reg-1 KO B7-H3-CAR T cells as a promising cell product for early-phase clinical testing in patients with solid tumors.

Detection of the off-target effects of base editors is important for identifying their safety risks but current methods for understanding their global activities have limitations in terms of sensitivity or bias by computationally selecting a subset of sites for experimental analysis. We present CHANGE-seq-BE, a method to assess the guide RNA-dependent off-target profile of both adenine and cytosine base editors that is simultaneously sensitive and unbiased. CHANGE-seq-BE relies on selective sequencing of base-editor-modified genomic DNA in vitro and provides comprehensive identification of genome-wide off-target mutations. We found that 98.8% of validated off-target sites were unique to ABE8e adenine base editors compared to Cas9 nuclease, suggesting substantially higher off-target activity of the former. We further applied CHANGE-seq-BE to support genotoxicity studies in an emergency investigational new drug application for customized adenine base editor treatment for a person with CD40L-deficient X-linked hyper IgM syndrome. Our results emphasize the importance of using a base-editor-specific method for identifying off-target activity.

Characterizing the protospacer adjacent motif (PAM) requirements of different Cas enzymes is a bottleneck in the discovery of Cas proteins and their engineered variants in mammalian cell contexts. Here, to overcome this challenge and to enable more scalable characterization of PAM preferences, we develop a method named GenomePAM that allows for direct PAM characterization in mammalian cells. GenomePAM leverages genomic repetitive sequences as target sites and does not require protein purification or synthetic oligos. GenomePAM uses a 20-nt protospacer that occurs ~16,942 times in every human diploid cell and is flanked by nearly random sequences. We demonstrate that GenomePAM can accurately characterize the PAM requirement of type II and type V nucleases, including the minimal PAM requirement of the near-PAMless SpRY and extended PAM for CjCas9. Beyond PAM characterization, GenomePAM allows for simultaneous comparison of activities and fidelities among different Cas nucleases on thousands of match and mismatch sites across the genome using a single gRNA and provides insight into the genome-wide chromatin accessibility profiles in different cell types.

CRISPR genetic therapies are revolutionizing the landscape of preclinical research and clinical studies, providing new potential routes for curative intervention for a range of previously untreatable diseases. As with any therapy, the therapeutic benefits and risks must be weighed against consideration of the disease threat. Genome-related adverse events are an inherent risk of CRISPR genetic therapies, including off-target edits. The perception that CRISPR therapies ought to have near-zero off-targets belies clinical medicine, therapy development and biology, which demonstrate that 'perfect' therapeutics do not exist. Given that not all genomic off-target events are equal, we provide a practical framework to evaluate and assess off-target safety based on the tools available today and ones that will be developed in the future. With the comprehensive information and assessment gathered using these guidelines, we aim to streamline the transition of CRISPR therapeutics from bench to bedside.

Genome editing enzymes can introduce targeted changes to the DNA in living cells 1-4 , transforming biological research and enabling the first approved gene editing therapy for sickle cell disease 5 . However, their genome-wide activity can be altered by genetic variation at on- or off-target sites 6-8 , potentially impacting both their precision and therapeutic safety. Due to a lack of scalable methods to measure genome-wide editing activity in cells from large populations and diverse target libraries, the frequency and extent of these variant effects on editing remains unknown. Here, we present the first population-scale study of how genetic variation affects the cellular genome-wide activity of CRISPR-Cas9, enabled by a novel, sensitive, and unbiased cellular assay, GUIDE-seq-2 with improved scalability and accuracy compared to the original broadly adopted method 9 . Analyzing Cas9 genome-wide activity at 1,115 on- and off-target sites across six guide RNAs in cells from 95 individuals spanning four genetically diverse populations, we found that variants frequently overlap off-target sites, with 13% significantly altering Cas9 editing activity by up to 33% indels. To understand common features of high-impact variants, we developed a new massively parallel biochemical assay, CHANGE-seq-R, to measure Cas9 activity across millions of mismatched target sites, and trained a deep neural network model, CHANGE-net, to accurately predict and interpret the effects of single-nucleotide variants on off-targets with up to six mismatches. Taken together, our findings illuminate a path to account for genetic variation when designing genome editing strategies for research and therapeutics.

Natural killer (NK) cells are an appealing off-the-shelf, allogeneic cellular therapy due to their cytotoxic profile. However, their activity against solid tumors remains suboptimal in part due to the upregulation of NK-inhibitory ligands, such as HLA-E, within the tumor microenvironment. Here, we utilize CRISPR-Cas9 to disrupt the KLRC1 gene (encoding the HLA-E-binding NKG2A receptor) and perform non-viral insertion of a GD2-targeting chimeric antigen receptor (CAR) within NK cells isolated from human peripheral blood. Genome editing with CRISPR/Cas9 ribonucleoprotein complexes yields efficient genomic disruption of the KLRC1 gene with 98% knockout efficiency and specific knock-in of the GD2 CAR transgene as high as 23%, with minimal off-target activity as shown by CHANGE-Seq, in-out PCR, amplicon sequencing and long read whole genome sequencing. KLRC1-GD2 CAR NK cells display high viability and proliferation, as well as precise cellular targeting and potency against GD2+ human tumor cells. Notably, KLRC1-GD2 CAR NK cells overcome HLA-E-based inhibition in vitro against HLA-E-expressing, GD2+ melanoma cells. Using a single-step, virus-free genome editing workflow, this study demonstrates the feasibility of precisely disrupting inhibitory signaling within NK cells via CRISPR/Cas9 while expressing a CAR to generate potent allogeneic cell therapies against HLA-E+ solid tumors.

Engineering and characterizing proteins can be time-consuming and cumbersome, motivating the development of generalist CRISPR-Cas enzymes1-4 to enable diverse genome-editing applications. However, such enzymes have caveats such as an increased risk of off-target editing3,5,6. Here, to enable scalable reprogramming of Cas9 enzymes, we combined high-throughput protein engineering with machine learning to derive bespoke editors that are more uniquely suited to specific targets. Through structure-function-informed saturation mutagenesis and bacterial selections, we obtained nearly 1,000 engineered SpCas9 enzymes and characterized their protospacer-adjacent motif (PAM)7 requirements to train a neural network that relates amino acid sequence to PAM specificity. By utilizing the resulting PAM machine learning algorithm (PAMmla) to predict the PAMs of 64 million SpCas9 enzymes, we identified efficacious and specific enzymes that outperform evolution-based and engineered SpCas9 enzymes as nucleases and base editors in human cells while reducing off-targets. An in silico-directed evolution method enables user-directed Cas9 enzyme design, including for allele-selective targeting of the RHOP23H allele in human cells and mice. Together, PAMmla integrates machine learning and protein engineering to curate a catalogue of SpCas9 enzymes with distinct PAM requirements, motivating a shift away from generalist enzymes towards safe and efficient bespoke Cas9 variants.

Brain-wide genome-editing remains a major hurdle for the treatment of neurogenetic disorders. Here, we report a non-viral, non-nanoparticle, chemical modification-based method, called Stimuli-Responsive Traceless Engineering Platform (STEP) that achieves highly efficient and brain-wide genome-editing of neurons in mice. Using cholesterol-based STEP as a lead, we show that a single administration of STEP-ribonucleoproteins (RNPs) results in functional genetic rescue with significant improvements across a battery of neurobehavioral domains in the Angelman syndrome (AS) mouse model. No significant off-target events or general toxicity effects are observed. STEP-RNPs are also highly efficient at editing human neurons and cortical brain organoids differentiated from AS patient-derived iPSCs. scRNA-seq analysis confirms functional genetic rescue via reactivation of Ube3a/UBE3A expression in human and mouse STEP-RNP-treated neuronal cells. Genome editing via STEP-RNPs has broad applications and the potential to treat many other neurogenetic disorders.

Pathogenic missense mutations in the alpha actin isotype 2 (ACTA2) gene cause multisystemic smooth muscle dysfunction syndrome (MSMDS), a genetic vasculopathy that is associated with stroke, aortic dissection and death in childhood. Here we perform mutation-specific protein engineering to develop a bespoke CRISPR–Cas9 enzyme with enhanced on-target activity against the most common MSMDS-causative mutation ACTA2 R179H. To directly correct the R179H mutation, we screened dozens of configurations of base editors to develop a highly precise corrective A-to-G edit with minimal deleterious bystander editing that is otherwise prevalent when using wild-type SpCas9 base editors. We create a murine model of MSMDS that shows phenotypes consistent with human patients, including vasculopathy and premature death, to explore the in vivo therapeutic potential of this strategy. Delivery of the customized base editor via an engineered smooth muscle-tropic adeno-associated virus (AAV-PR) vector substantially prolongs survival and rescues systemic phenotypes across the lifespan of MSMDS mice, including in the vasculature, aorta and brain. Our results highlight how bespoke mutant-specific CRISPR–Cas9 enzymes can improve mutation correction with base editors. Engineering a mutant-specific customized base editor precisely corrects a mutation while minimizing bystander edits, leading to substantial phenotypic recovery in mouse models of multisystemic smooth muscle dysfunction syndrome.

Editing the +58 region of the BCL11A erythroid enhancer has shown promise in treating β-globin disorders. To address variations in fetal hemoglobin (HbF) response, we investigated editing both +58 and +55 enhancers. Rhesus macaques transplanted with edited hematopoietic stem/progenitor cells (HSPCs) following busulfan conditioning exhibited durable, high-level (∼90%) editing frequencies post transplantation with sustained HbF reactivation over 4 years, without hematological perturbations. HbF levels were further boosted by stress erythropoiesis or hydroxyurea. Bone marrow analysis revealed that gene edits were predominantly programmed deletions, programmed inversions, and short indels, each disrupting the enhancer core TGN7-9WGATAR half E-box/GATA binding motifs. Nonprogrammed long deletions were disfavored in engrafting cells. CD45 antibody-drug conjugate (ADC) conditioning achieved comparable engraftment and HbF reactivation, whereas lentiviral vector tracking showed polyclonal reconstitution with dynamics similar to animals conditioned with total body irradiation (TBI) or busulfan. Joining CD45-ADC conditioning with combined enhancer editing presents an effective strategy for β-hemoglobinopathies, enabling durable HbF reactivation without chemotherapy.

Genetic vascular disorders are prevalent diseases that have diverse etiologies and few treatment options. Pathogenic missense mutations in the alpha actin isotype 2 gene (ACTA2) primarily affect smooth muscle cell (SMC) function and cause multisystemic smooth muscle dysfunction syndrome (MSMDS), a genetic vasculopathy that is associated with stroke, aortic dissection, and death in childhood. Here, we explored genome editing to correct the most common MSMDS-causative mutation ACTA2 R179H. In a first-in-kind approach, we performed mutation-specific protein engineering to develop a bespoke CRISPR-Cas9 enzyme with enhanced on-target activity against the R179H sequence. To directly correct the R179H mutation, we screened dozens of configurations of base editors (comprised of Cas9 enzymes, deaminases, and gRNAs) to develop a highly precise corrective A-to-G edit with minimal deleterious bystander editing that is otherwise prevalent when using wild-type SpCas9 base editors. We then created a murine model of MSMDS that exhibits phenotypes consistent with human patients, including vasculopathy and premature death, to explore the in vivo therapeutic potential of this base editing strategy. Delivery of the customized base editor via an engineered SMC-tropic adeno-associated virus (AAV-PR) vector substantially prolonged survival and rescued systemic phenotypes across the lifespan of MSMDS mice, including in the vasculature, aorta, and brain. Together, our optimization of a customized base editor highlights how bespoke CRISPR-Cas enzymes can enhance on-target correction while minimizing bystander edits, culminating in a precise editing approach that may enable a long-lasting treatment for patients with MSMDS.

Sickle cell disease (SCD) is a common, severe genetic blood disorder. Current pharmacotherapies are partially effective and allogeneic hematopoietic stem cell transplantation is associated with immune toxicities. Genome editing of patient hematopoietic stem cells (HSCs) to reactivate fetal hemoglobin (HbF) in erythroid progeny offers an alternative potentially curative approach to treat SCD. Although the FDA released guidelines for evaluating genome editing risks, it remains unclear how best to approach pre-clinical assessment of genome-edited cell products. Here, we describe rigorous pre-clinical development of a therapeutic γ-globin gene promoter editing strategy that supported an investigational new drug application cleared by the FDA. We compared γ-globin promoter and BCL11A enhancer targets, identified a potent HbF-inducing lead candidate, and tested our approach in mobilized CD34+ hematopoietic stem progenitor cells (HSPCs) from SCD patients. We observed efficient editing, HbF induction to predicted therapeutic levels, and reduced sickling. With single-cell analyses, we defined the heterogeneity of HbF induction and HBG1/HBG2 transcription. With CHANGE-seq for sensitive and unbiased off-target discovery followed by targeted sequencing, we did not detect off-target activity in edited HSPCs. Our study provides a blueprint for translating new ex vivo HSC genome editing strategies toward clinical trials for treating SCD and other blood disorders.

Background: Sickle cell disease (SCD) symptoms can be alleviated with elevated expression of fetal hemoglobin (HbF) in red blood cells (RBCs). CRISPR-Cas9 editing to produce indels disrupting DNA regulatory elements that repress g-globin gene expression can induce HbF in adult RBCs. We have previously shown that the disruption of the BCL11A repressor-binding motifs (-115) in the g-globin gene promoters can effectively induce HbF that mimics naturally occurring hereditary persistent fetal hemoglobin variants. Here we present our clinical scale up and optimization, pharmacological, and toxicological studies in support of our recently approved FDA IND application to initiate our St. Jude Autologous Genome Edited Stem Cell (SAGES1) clinical trial (NCT06506461). Methods: To translate this editing strategy for clinical application, we optimized RNP electroporation of human donor CD34+ HSPCs with Cas9 and the HBG-115 gRNA. For maximal editing rates and cell recovery, we explored the Cas9 concentration, RNP ratio, and electroporation cell concentration using reagents produced in a Good Manufacturing Practice (GMP) facility. We then scaled up the electroporation protocol to the clinically relevant MaxCyte electroporator. Several other aspects of the culture and electroporation protocol were then investigated to establish optimal conditions and acceptable ranges for an efficient and robust process, including prestimulation cell culture, cryopreservation, and RNP complexation. Pre-GMP engineering runs were conducted using full scale healthy donor plerixafor mobilized HSPCs (n=3) edited with GMP-like Cas9 protein and sgRNA. The Cas9/HBG-115 edited cells were xenotransplanted into NBSGW mice for both in vivo pharmacological and in vivo toxicological studies. Results: With our scaled and optimized ex vivo editing protocol, we demonstrated that the Cas9/HBG-115 editing rate of CD34+ HSPCs and HbF expression correlates directly with increasing Cas9 concentration, and that at optimal concentrations, we can achieve >80% editing and ~25% HbF expression in erythroid cells in vitro. In our in vivo pharmacological study, we obtained ~92% editing in the bulk HSPCs before xenotransplantation. Edited and unedited cells engrafted efficiently. We retained high indel rates (70-84%) from bone marrow harvested after 16 weeks. We attained HbF levels of 17.1-25.9% from erythroid cells isolated from the mice bone marrow. We observed no differences in human engraftment or lineages in transplanted mice compared to unedited controls. Indel pattern distribution and HSPC subpopulation frequency of edited CD34+ HSPCs were similar for in vitro and engrafted samples within the same donor. Population frequencies of human hematopoietic lineages were similar between edited and unedited cells and editing rates between human lineages of edited CD34+ HSPCs were also similar to bulk editing rates. From our in vivo toxicological studies, we also did not detect any adverse effects in the mice and there was no detectable off target editing. Conclusion: Our IND package demonstrating an optimized, safe, and effective preclinical treatment of Cas9/HBG-115 edited CD34+ HSPCs was approved by the FDA to proceed with the clinical trial that will begin accrual soon.

Spinal muscular atrophy (SMA) is caused by mutations in SMN1 . SMN2 is a paralogous gene with a C•G-to-T•A transition in exon 7, which causes this exon to be skipped in most SMN2 transcripts, and results in low levels of the protein survival motor neuron (SMN). Here we show, in fibroblasts derived from patients with SMA and in a mouse model of SMA that, irrespective of the mutations in SMN1 , adenosine base editors can be optimized to target the SMN2 exon-7 mutation or nearby regulatory elements to restore the normal expression of SMN. After optimizing and testing more than 100 guide RNAs and base editors, and leveraging Cas9 variants with high editing fidelity that are tolerant of different protospacer-adjacent motifs, we achieved the reversion of the exon-7 mutation via an A•T-to-G•C edit in up to 99% of fibroblasts, with concomitant increases in the levels of the SMN2 exon-7 transcript and of SMN . Targeting the SMN2 exon-7 mutation via base editing or other CRISPR-based methods may provide long-lasting outcomes to patients with SMA.

Haematopoietic stem cell (HSC) transplantation (HSCT) is the only curative treatment for a broad range of haematological malignancies, but the standard of care relies on untargeted chemotherapies and limited possibilities to treat malignant cells after HSCT without affecting the transplanted healthy cells1. Antigen-specific cell-depleting therapies hold the promise of much more targeted elimination of diseased cells, as witnessed in the past decade by the revolution of clinical practice for B cell malignancies2. However, target selection is complex and limited to antigens expressed on subsets of haematopoietic cells, resulting in a fragmented therapy landscape with high development costs2-5. Here we demonstrate that an antibody-drug conjugate (ADC) targeting the pan-haematopoietic marker CD45 enables the antigen-specific depletion of the entire haematopoietic system, including HSCs. Pairing this ADC with the transplantation of human HSCs engineered to be shielded from the CD45-targeting ADC enables the selective eradication of leukaemic cells with preserved haematopoiesis. The combination of CD45-targeting ADCs and engineered HSCs creates an almost universal strategy to replace a diseased haematopoietic system, irrespective of disease aetiology or originating cell type. We propose that this approach could have broad implications beyond haematological malignancies.

Staphylococcus aureus Cas9 (SaCas9), which is smaller than the widely-used Streptococcus pyogenes Cas9 (SpCas9), has been harnessed for gene therapy using an adeno-associated virus vector. However, SaCas9 requires an NNGRRT (where N is any nucleotide and R is A or G) protospacer adjacent motif (PAM) for target DNA recognition, thereby restricting the targeting range. In addition, the nuclease activation mechanism of SaCas9 remains elusive. Here, we rationally engineered a SaCas9 variant (eSaCas9-NNG) with an expanded target scope and reduced off-target activity. eSaCas9-NNG induced indels and base conversions at endogenous sites bearing NNG PAMs in human cells and mice. We further determined the cryo-electron microscopy structures of eSaCas9-NNG in four sequential states, PAM-checking state, DNA-unwinding state, pre-catalytic state and catalytically active state, which illuminate notable differences in the activation mechanisms between small SaCas9 and larger SpCas9. Overall, our findings demonstrate that eSaCas9-NNG could be used as a versatile genome editing tool for in vivo gene therapy, and improve our mechanistic understanding of the diverse CRISPR-Cas9 nucleases.