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Dominant missense pathogenic variants in cardiac myosin heavy chain cause hypertrophic cardiomyopathy (HCM), a currently incurable disorder that increases risk for stroke, heart failure and sudden cardiac death. In this study, we assessed two different genetic therapies—an adenine base editor (ABE8e) and a potent Cas9 nuclease delivered by AAV9—to prevent disease in mice carrying the heterozygous HCM pathogenic variant myosin R403Q. One dose of dual-AAV9 vectors, each carrying one half of RNA-guided ABE8e, corrected the pathogenic variant in ≥70% of ventricular cardiomyocytes and maintained durable, normal cardiac structure and function. An additional dose provided more editing in the atria but also increased bystander editing. AAV9 delivery of RNA-guided Cas9 nuclease effectively inactivated the pathogenic allele, albeit with dose-dependent toxicities, necessitating a narrow therapeutic window to maintain health. These preclinical studies demonstrate considerable potential for single-dose genetic therapies to correct or silence pathogenic variants and prevent the development of HCM.

Although cyclophilins are attractive targets for probing biology and therapeutic intervention, no subtype-selective cyclophilin inhibitors have been described. We discovered novel cyclophilin inhibitors from the in vitro selection of a DNA-templated library of 256,000 drug-like macrocycles for cyclophilin D (CypD) affinity. Iterated macrocycle engineering guided by ten X-ray co-crystal structures yielded potent and selective inhibitors (half maximal inhibitory concentration (IC50) = 10 nM) that bind the active site of CypD and also make novel interactions with non-conserved residues in the S2 pocket, an adjacent exo-site. The resulting macrocycles inhibit CypD activity with 21- to >10,000-fold selectivity over other cyclophilins and inhibit mitochondrial permeability transition pore opening in isolated mitochondria. We further exploited S2 pocket interactions to develop the first cyclophilin E (CypE)-selective inhibitor, which forms a reversible covalent bond with a CypE S2 pocket lysine, and exhibits 30- to >4,000-fold selectivity over other cyclophilins. These findings reveal a strategy to generate isoform-selective small-molecule cyclophilin modulators, advancing their suitability as targets for biological investigation and therapeutic development.

Despite the availability of Cas9 variants with varied protospacer-adjacent motif (PAM) compatibilities, some genomic loci—especially those with pyrimidine-rich PAM sequences—remain inaccessible by high-activity Cas9 proteins. Moreover, broadening PAM sequence compatibility through engineering can increase off-target activity. With directed evolution, we generated four Cas9 variants that together enable targeting of most pyrimidine-rich PAM sequences in the human genome. Using phage-assisted noncontinuous evolution and eVOLVER-supported phage-assisted continuous evolution, we evolved Nme2Cas9, a compact Cas9 variant, into variants that recognize single-nucleotide pyrimidine-PAM sequences. We developed a general selection strategy that requires functional editing with fully specified target protospacers and PAMs. We applied this selection to evolve high-activity variants eNme2-T.1, eNme2-T.2, eNme2-C and eNme2-C.NR. Variants eNme2-T.1 and eNme2-T.2 offer access to N4TN PAM sequences with comparable editing efficiencies as existing variants, while eNme2-C and eNme2-C.NR offer less restrictive PAM requirements, comparable or higher activity in a variety of human cell types and lower off-target activity at N4CN PAM sequences.

The viral delivery of base editors has been complicated by their size and by the limited packaging capacity of adeno-associated viruses (AAVs). Typically, dual-AAV approaches based on trans-splicing inteins have been used. Here we show that, compared with dual-AAV systems, AAVs with size-optimized genomes incorporating compact adenine base editors (ABEs) enable efficient editing in mice at similar or lower doses. Single-AAV-encoded ABEs retro-orbitally injected in mice led to editing efficiencies in liver (66%), heart (33%) and muscle (22%) tissues that were up to 2.5-fold those of dual-AAV ABE8e, and to a 93% knockdown (on average) of human PCSK9 and of mouse Pcsk9 and Angptl3 in circulation, concomitant with substantial reductions of plasma cholesterol and triglycerides. Moreover, three size-minimized ABE8e variants, each compatible with single-AAV delivery, collectively offer compatibility with protospacer-adjacent motifs for editing approximately 82% of the adenines in the human genome. ABEs encoded within single AAVs will facilitate research and therapeutic applications of base editing by simplifying AAV production and characterization, and by reducing the dose required for the desired level of editing.

Realizing the promise of prime editing for the study and treatment of genetic disorders requires efficient methods for delivering prime editors (PEs) in vivo. Here we describe the identification of bottlenecks limiting adeno-associated virus (AAV)-mediated prime editing in vivo and the development of AAV-PE vectors with increased PE expression, prime editing guide RNA stability and modulation of DNA repair. The resulting dual-AAV systems, v1em and v3em PE-AAV, enable therapeutically relevant prime editing in mouse brain (up to 42% efficiency in cortex), liver (up to 46%) and heart (up to 11%). We apply these systems to install putative protective mutations in vivo for Alzheimer’s disease in astrocytes and for coronary artery disease in hepatocytes. In vivo prime editing with v3em PE-AAV caused no detectable off-target effects or significant changes in liver enzymes or histology. Optimized PE-AAV systems support the highest unenriched levels of in vivo prime editing reported to date, facilitating the study and potential treatment of diseases with a genetic component.

Spinal muscular atrophy (SMA), the leading genetic cause of infant mortality, arises from survival motor neuron (SMN) protein insufficiency resulting from SMN1 loss. Approved therapies circumvent endogenous SMN regulation and require repeated dosing or may wane. We describe genome editing of SMN2 , an insufficient copy of SMN1 harboring a C6>T mutation, to permanently restore SMN protein levels and rescue SMA phenotypes. We used nucleases or base editors to modify five SMN2 regulatory regions. Base editing converted SMN2 T6>C, restoring SMN protein levels to wild type. Adeno-associated virus serotype 9–mediated base editor delivery in Δ7SMA mice yielded 87% average T6>C conversion, improved motor function, and extended average life span, which was enhanced by one-time base editor and nusinersen coadministration (111 versus 17 days untreated). These findings demonstrate the potential of a one-time base editing treatment for SMA.

Guide RNAs offer programmability for CRISPR-Cas9 genome editing but also add challenges for delivery. Chemical modification, which has been key to the success of oligonucleotide therapeutics, can enhance the stability, distribution, cellular uptake, and safety of nucleic acids. Previously, we engineered heavily and fully modified SpyCas9 crRNA and tracrRNA, which showed enhanced stability and retained activity when delivered to cultured cells in the form of the ribonucleoprotein complex. In this study, we report that a short, fully stabilized oligonucleotide (a “protecting oligo”), which can be displaced by tracrRNA annealing, can significantly enhance the potency and stability of a heavily modified crRNA. Furthermore, protecting oligos allow various bioconjugates to be appended, thereby improving cellular uptake and biodistribution of crRNAin vivo. Finally, we achievedin vivogenome editing in adult mouse liver and central nervous system via co-delivery of unformulated, chemically modified crRNAs with protecting oligos and AAV vectors that express tracrRNA and either SpyCas9 or a base editor derivative. Our proof-of-concept establishment of AAV/crRNA co-delivery offers a route towards transient editing activity, target multiplexing, guide redosing, and vector inactivation.

Cas9 is a programmable nuclease that has furnished transformative technologies, including base editors and transcription modulators (e.g., CRISPRi/a), but several applications of these technologies, including therapeutics, mandatorily require precision control of their half-life. For example, such control can help avert any potential immunological and adverse events in clinical trials. Current genome editing technologies to control the half-life of Cas9 are slow, have lower activity, involve fusion of large response elements (> 230 amino acids), utilize expensive controllers with poor pharmacological attributes, and cannot be implementedin vivoon several CRISPR-based technologies. We report a general platform for half-life control using the molecular glue, pomalidomide, that binds to a ubiquitin ligase complex and a response-element bearing CRISPR-based technology, thereby causing the latter’s rapid ubiquitination and degradation. Using pomalidomide, we were able to control the half-life of large CRISPR-based technologies (e.g., base editors, CRISPRi) and small anti-CRISPRs that inhibit such technologies, allowing us to build the first examples of on-switch for base editors. The ability to switch on, fine-tune and switch-off CRISPR-based technologies with pomalidomide allowed complete control over their activity, specificity, and genome editing outcome. Importantly, the miniature size of the response element and favorable pharmacological attributes of the drug pomalidomide allowed control of activity of base editorin vivousing AAV as the delivery vehicle. These studies provide methods and reagents to precisely control the dosage and half-life of CRISPR-based technologies, propelling their therapeutic development.

The all-protein cytosine base editor DdCBE uses TALE proteins and a double-stranded DNA-specific cytidine deaminase (DddA) to mediate targeted C•G-to-T•A editing. To improve editing efficiency and overcome the strict TC sequence-context constraint of DddA, we used phage-assisted non-continuous and continuous evolution to evolve DddA variants with improved activity and expanded targeting scope. Compared to canonical DdCBEs, base editors with evolved DddA6 improved mitochondrial DNA (mtDNA) editing efficiencies at TC by 3.3-fold on average. DdCBEs containing evolved DddA11 offered a broadened HC (H = A, C or T) sequence compatibility for both mitochondrial and nuclear base editing, increasing average editing efficiencies at AC and CC targets from less than 10% for canonical DdCBE to 15–30% and up to 50% in cell populations sorted to express both halves of DdCBE. We used these evolved DdCBEs to efficiently install disease-associated mtDNA mutations in human cells at non-TC target sites. DddA6 and DddA11 substantially increase the effectiveness and applicability of all-protein base editing.

On-target toxicity to normal cells is a major safety concern with targeted immune and gene therapies. Here, we developed a base editing (BE) approach exploiting a naturally occurring CD33 single nucleotide polymorphism leading to removal of full-length CD33 surface expression on edited cells. CD33 editing in human and nonhuman primate (NHP) hematopoietic stem and progenitor cells (HSPCs) protects from CD33-targeted therapeutics without affecting normal hematopoiesisin vivo, thus demonstrating potential for novel immunotherapies with reduced off-leukemia toxicity. For broader applications to gene therapies, we demonstrated highly efficient (>70%) multiplexed adenine base editing of the CD33 and gamma globin genes, resulting in long-term persistence of dual gene-edited cells with HbF reactivation in NHPs.In vitro, dual gene-edited cells could be enriched via treatment with the CD33 antibody-drug conjugate, gemtuzumab ozogamicin (GO). Together, our results highlight the potential of adenine base editors for improved immune and gene therapies.Graphical abstract

Sickle Cell Disease (SCD) is a monogenic disease caused by a nucleotide mutation in the β-globin gene. Current gene therapy studies are mainly focused on lentivirus vector-mediated gene addition or CRISPR/Cas9-mediated fetal globin reactivation, leaving the root cause unfixed. We developed a vectorized prime editing system that can directly repair the SCD mutation in hematopoietic stem cells (HSCs) in vivo in a SCD mouse model (CD46/Townes mice). Our approach involved a single intravenous injection of a non-integrating, prime editor-expressing virus vector into mobilized CD46/Townes mice and low-dose drug selection in vivo. This procedure resulted in the correction of ~40% of bS alleles in HSCs. On average 43% of HbS was replaced by HbA thereby greatly mitigating the SCD phenotypes. Transplantation in secondary recipients demonstrated that long-term repopulating HSCs were edited. Highly efficient target site editing was achieved with minimal generation of insertions and deletions and no detectable off-target editing. Because of its simplicity and portability, our in vivo prime editing approach has the potential for application in resource-poor countries where SCD is prevalent.

Prime editing enables the installation of virtually any combination of point mutations, small insertions or small deletions in the DNA of living cells. A prime editing guide RNA (pegRNA) directs the prime editor protein to the targeted locus and also encodes the desired edit. Here we show that degradation of the 3′ region of the pegRNA that contains the reverse transcriptase template and the primer binding site can poison the activity of prime editing systems, impeding editing efficiency. We incorporated structured RNA motifs to the 3′ terminus of pegRNAs that enhance their stability and prevent degradation of the 3′ extension. The resulting engineered pegRNAs (epegRNAs) improve prime editing efficiency 3–4-fold in HeLa, U2OS and K562 cells and in primary human fibroblasts without increasing off-target editing activity. We optimized the choice of 3′ structural motif and developed pegLIT, a computational tool to identify non-interfering nucleotide linkers between pegRNAs and 3′ motifs. Finally, we showed that epegRNAs enhance the efficiency of the installation or correction of disease-relevant mutations. Stabilizing pegRNAs with 3′ RNA structures increases prime editing efficiency.

Mutations in the CFTR gene that lead to premature stop codons or splicing defects cause cystic fibrosis (CF) and are not amenable to treatment by small-molecule modulators. Here, we investigate the use of adenine base editor (ABE) ribonucleoproteins (RNPs) that convert A•T to G•C base pairs as a therapeutic strategy for three CF-causing mutations. Using ABE RNPs, we corrected in human airway epithelial cells premature stop codon mutations (R553X and W1282X) and a splice-site mutation (3849 + 10 kb C > T). Following ABE delivery, DNA sequencing revealed correction of these pathogenic mutations at efficiencies that reached 38–82% with minimal bystander edits or indels. This range of editing was sufficient to attain functional correction of CFTR-dependent anion channel activity in primary epithelial cells from CF patients and in a CF patient-derived cell line. These results demonstrate the utility of base editor RNPs to repair CFTR mutations that are not currently treatable with approved therapeutics.

Chemical probes are lacking for most human proteins. Covalent chemistry represents an attractive strategy for expanding the ligandability of the proteome, and chemical proteomics has revealed numerous electrophile-reactive cysteines on diverse proteins. Determining which of these covalent binding events impact protein function, however, remains challenging. Here, we describe a base-editing strategy to infer the functionality of cysteines by quantifying the impact of their missense mutation on cell proliferation. We show that the resulting atlas, which covers >13,800 cysteines on >1,750 cancer dependency proteins, correctly predicts the essentiality of cysteines targeted by cancer therapeutics and, when integrated with chemical proteomic data, identifies essential, ligandable cysteines on >110 cancer dependency proteins. We further demonstrate how measurements of reactivity in native versus denatured proteomes can discriminate essential cysteines amenable to chemical modification from those buried in protein structures, providing a valuable resource to prioritize the pursuit of small-molecule probes with high function-perturbing potential.

CRISPR-Cas-based genome editing technologies could, in principle, be used to treat a wide variety of inherited diseases, including genetic disorders of vision. Programmable CRISPR-Cas nucleases are effective tools for gene disruption, but they are poorly suited for precisely correcting pathogenic mutations in most therapeutic settings. Recently developed precision genome editing agents, including base editors and prime editors, have enabled precise gene correction and disease rescue in multiple preclinical models of genetic disorders. Additionally, new delivery technologies that transiently deliver precision genome editing agents in vivo offer minimized off-target editing and improved safety profiles. These improvements to precision genome editing and delivery technologies are expected to revolutionize the treatment of genetic disorders of vision and other diseases. In this Perspective, we describe current preclinical and clinical genome editing approaches for treating inherited retinal degenerative diseases, and we discuss important considerations that should be addressed as these approaches are translated into clinical practice.

Programmable C•G-to-G•C base editors (CGBEs) have broad scientific and therapeutic potential, but their editing outcomes have proved difficult to predict and their editing efficiency and product purity are often low. We describe a suite of engineered CGBEs paired with machine learning models to enable efficient, high-purity C•G-to-G•C base editing. We performed a CRISPR interference (CRISPRi) screen targeting DNA repair genes to identify factors that affect C•G-to-G•C editing outcomes and used these insights to develop CGBEs with diverse editing profiles. We characterized ten promising CGBEs on a library of 10,638 genomically integrated target sites in mammalian cells and trained machine learning models that accurately predict the purity and yield of editing outcomes (R = 0.90) using these data. These CGBEs enable correction to the wild-type coding sequence of 546 disease-related transversion single-nucleotide variants (SNVs) with >90% precision (mean 96%) and up to 70% efficiency (mean 14%). Computational prediction of optimal CGBE-single-guide RNA pairs enables high-purity transversion base editing at over fourfold more target sites than achieved using any single CGBE variant.

Sickle cell disease (SCD) is caused by a mutation in the β-globin gene HBB1. We used a custom adenine base editor (ABE8e-NRCH)2,3 to convert the SCD allele (HBBS) into Makassar β-globin (HBBG), a non-pathogenic variant4,5. Ex vivo delivery of mRNA encoding the base editor with a targeting guide RNA into haematopoietic stem and progenitor cells (HSPCs) from patients with SCD resulted in 80% conversion of HBBS to HBBG. Sixteen weeks after transplantation of edited human HSPCs into immunodeficient mice, the frequency of HBBG was 68% and hypoxia-induced sickling of bone marrow reticulocytes had decreased fivefold, indicating durable gene editing. To assess the physiological effects of HBBS base editing, we delivered ABE8e-NRCH and guide RNA into HSPCs from a humanized SCD mouse6 and then transplanted these cells into irradiated mice. After sixteen weeks, Makassar β-globin represented 79% of β-globin protein in blood, and hypoxia-induced sickling was reduced threefold. Mice that received base-edited HSPCs showed near-normal haematological parameters and reduced splenic pathology compared to mice that received unedited cells. Secondary transplantation of edited bone marrow confirmed that the gene editing was durable in long-term haematopoietic stem cells and showed that HBBS-to-HBBG editing of 20% or more is sufficient for phenotypic rescue. Base editing of human HSPCs avoided the p53 activation and larger deletions that have been observed following Cas9 nuclease treatment. These findings point towards a one-time autologous treatment for SCD that eliminates pathogenic HBBS, generates benign HBBG, and minimizes the undesired consequences of double-strand DNA breaks. A custom adenine base editor can edit the variant of the β-globin gene that causes sickle cell disease into a non-pathogenic variant in human and mouse cells, and transplantation of the edited cells rescues sickle cell disease in mice.

Leber Congenital Amaurosis (LCA16) is a progressive vision loss disorder caused by point mutations in the KCNJ13 gene, which encodes an inward-rectifying potassium channel, Kir7.1. A nonsense mutation, W53X (c.158G>A), leads to premature truncation of the protein, which disrupts K+ conductance and makes retinal pigmented epithelium (RPE) non-functional. While there are no current treatments for LCA16, here we explore CRISPR base editing as a strategy to correct the single base pair pathogenic variant. We show that silica nanoparticle (SNC)-mediated delivery of an adenine base editor (ABE8e) mRNA and single-guide RNA can precisely and efficiently correct the KCNJ13W53X mutation to restore Kir7.1 channel function in an induced pluripotent stem cell-derived RPE (iPSC-RPE) model of LCA16, as well as in patient-derived fibroblasts and in a new LCA16 mouse model. We observed a higher editing efficiency in LCA16 patient-derived fibroblasts (47.38% ± 1.02) than in iPSC-RPE (16.90% ± 1.58) with no detectable off-target editing. The restored channel function in the edited cells resulted in a phenotype comparable to wild-type cells, suggesting the possibility that this treatment could restore vision in LCA16 patients. Injection of ABE8e-carrying SNCs decorated with all-trans retinoic acid (ATRA) ligand into the subretinal space in LCA16 mice showed specific delivery only to RPE and resulted in gene correction in approximately 10% of RPE cells that received the editing machinery. We observed marginal recovery of electroretinogram (ERG) following correction of the W53X allele at the injection site with no further degeneration of RPE. These findings provide a foundation and a proof-of-concept to transition from bench to bedside and support its further development for treating pediatric blindness using a safer mode of SNC-mediated delivery of base editors.

Point mutations in the KCNJ13 gene cause an autosomal recessive, childhood blindness, Leber congenital amaurosis (LCA16) due to a loss-of-function Kir7.1 channel. In the present study, we investigated the etiology of LCA16 caused by a KCNJ13 missense mutation (c.431T>C, p.Leu144Pro) and explored the activity of two cytosine base editors mRNAs (CBEs, BE4max-WTCas9, and evoCDA-SpCas9-NG) as a proof-of-concept therapeutic option. We observed the KCNJ13-related retinopathy phenotype in patients harboring L144P mutation. Our in-silico prediction and in vitro validation demonstrated that L144P mutation affects the channel function. We observed high on-target efficiency in the CBEs treated L144P mutant gene expressing HEK-293 cells. Strikingly, our evaluation of base editing efficacy using electrophysiology showed negligible channel function. We found that the editing bystander ‘Cs’ in the protospacer region led to a missense change (L143F) in evoCDA edited cells and only silent changes in BE4max edited cells. Upon investigation of the effect of the synonymous codon, our extended analysis revealed distortion of mRNA structure, altered half-life, and/or low abundance of the cognate tRNA. We propose that KCNJ13-L144P mutation or other genes that share similar genetic complexity may be challenging to correct with the current generation of CRISPR base editors, and a combinational therapy using CRISPR base editors with a tighter editing window and requisite cognate-tRNA supplementation could be an alternative therapeutic approach to restore Kir7.1 channel function in LCA16 patients. Other options for hard-to-rescue alleles could employ homology-directed repair using CRISPR/Cas9 nucleases, Prime editing, and AAV-mediated gene augmentation.