Tumor-infiltrating lymphocyte (TIL) therapies harness tumor-specific T cells endogenous to a patient's repertoire but their efficacy is limited by challenges such as low frequencies of tumor-specific clonotypes and dysfunctional T cell phenotypes. These challenges necessitate technologies to engineer and reprogram endogenous tumor-specific TILs ex vivo. Here, we present a strategy using engineered virus-like particles (eVLPs) pseudotyped with peptide-major histocompatibility complexes (pMHCs) as a programmable, single-effector platform for selective and coordinated priming, expansion, and genome editing of rare antigen-specific CD8+ T cells among their endogenous polyclonal repertoires. We demonstrate that pMHC-pseudotyped eVLPs (pMHC-eVLPs) deliver T cell function-enhancing base editors to arm polyclonal lymphocytes with enhanced anti-tumor cytotoxicity by selectively expanding and engineering the tumor-specific T cell compartment. Our work establishes pMHC-eVLPs as a platform for enhancing TIL therapy with precision gene edits without the risks of bystander T cell engineering associated with polyclonal TIL engineering approaches.

Prime editing is a versatile clinical genome editing method that enables precise substitutions, small insertions and deletions at specified locations in the genomes of living systems including human cells. Although non-viral lipid nanoparticle (LNP) delivery of RNA in vivo has become a preferred method for gene editing in animals and patients, its application to complex, three-component prime editing systems has yielded low editing efficiencies. Here we developed a systematic prime editing LNP (PE-LNP) optimization platform that addresses key bottlenecks in cargo design that limit editing efficiency. This generalizable workflow yielded PE-LNPs that can achieve 49% average in vivo prime editing in the bulk mouse liver with a single dose of 2 mg kg-1. We applied our workflow to the correction of PAH R408W, a cause of phenylketonuria, in a mouse model and achieved prime editing efficiencies and serum phenylalanine levels anticipated to be curative. We also show that PE-LNPs minimize off-target editing compared with DNA delivery methods, induce only transient elevation of liver enzymes and can be dosed repeatedly to improve editing efficiencies. These PE-LNP systems provide an attractive alternative to viral delivery by offering transient expression that minimizes off-target editing, no observed long-term toxicity and high levels of non-viral in vivo liver prime editing.

Chemical proteomics has identified covalent ligands targeting cysteine residues across many hundreds of human proteins. The functional effects of these liganding events, however, remain challenging to assign at scale. Here we describe ESCAPE (Endogenous Site-specific Competition Assays using Prime Editors), a platform for the site-resolved functional analysis of covalent ligands in cells. In this method, cysteine-to-serine substitutions are generated by prime editing to abrogate covalent ligand-protein interactions, and the impact of these edits on ligand-induced cellular phenotypes is quantified through allele frequency-based resistance scores. Applied to ligandable cysteines mapped by activity-based protein profiling in 50+ proteins, ESCAPE identified multiple covalent ligand-protein interactions that impair cancer cell growth, including azetidine butynamides that target a non-orthosteric cysteine in the RNA helicase DDX49 to disrupt 18S rRNA processing, 40S ribosome assembly, and protein synthesis. ESCAPE thus provides a scalable framework for the functional characterization of covalent ligands targeting structurally and mechanistically diverse proteins.

Understanding how the first cell lineages in human development are specified and maintained has fundamental importance and clinical implications for regenerative medicine, infertility and pregnancy loss. Although mouse models have provided valuable insights into transcription factors regulating early development, translating these findings to human embryos has been limited by ethical, technical and biological constraints. Functional studies of transcription factors in human embryos have been hindered by nuclease-based genome editing approaches that induce genotoxicity1-3. Here, to overcome this, we applied ABE8e adenine base editing4,5 to precisely target an exon splice donor site, resulting in a splicing defect and functional knockout of the developmental regulator NANOG in human embryos. This approach did not trigger genotoxicity and showed limited off-target editing. Loss of NANOG disrupts pluripotent epiblast specification and instead cells differentiate towards a primitive endoderm (yolk sac) or trophectoderm (placental) transcriptional programme. Retention of primitive endoderm differentiation in NANOG-edited human embryos reveals a functional compensation that is distinct from mouse, underscoring the importance of directly investigating human development. Our findings demonstrate an essential role for NANOG in human pluripotency and epiblast specification and highlight the utility of base editing for functional interrogation of human development.

Primary human myeloid cells hold promise for immunotherapies, yet efficient, scalable technologies for engineering and screening in these cells remain limited. Here we present a virus-like particle (VLP)-based toolkit that delivers diverse CRISPR editing modalities to human monocytes, macrophages and dendritic cells with high efficiency while preserving viability and innate immune responsiveness. VLP-mediated delivery of ribonucleoproteins supports gene knockout, base editing and epigenetic silencing. Combined with adeno-associated virus-mediated donor delivery, this approach enables site-specific integration of large DNA sequences by homology-directed repair. We developed SLICeVLP, which pairs sgRNA delivery by VPX-lentivirus with Cas9 protein delivery by engineered VLPs, and used it for pooled loss-of-function and Perturb-seq screens in human macrophages. We uncovered regulators of tumor necrosis factor (TNF) and CD80 expression, converging on TNFAIP3 as a central regulator of inflammatory polarization. TNFAIP3 ablation drove a proinflammatory state resistant to suppressive repolarization and enhanced cytotoxicity in chimeric antigen receptor macrophages. This system enables unbiased functional genomics in primary human myeloid cells, with implications for myeloid cell therapy design.

CRISPR/Cas9-based gene-editing technologies offer promise for treating inherited retinal diseases (IRDs), however safe and efficient ocular delivery of precision editors remains challenging. To address this challenge, we report a class of Coomassie brilliant blue (CBB)-derived lipidoids that bind and deliver proteins. Subretinal injection of Cre complexed with these lipidoids into mT/mG mice leads to robust recombination in the retinal pigment epithelium and photoreceptors. We employ the CBB-lipidoid platform to deliver adenine base editor (ABE) ribonucleoproteins (RNP). Incorporating CBB lipidoids into liposomes improves delivery efficiency. CBB11 stands out for facilitating precise in vivo ABE-mediated gene editing. Delivery of liposome-CBB11-RNP complexes results in a 120-fold increase in base editing compared to RNP alone and restores the scotopic ERG b-wave response in the rd12 mouse model. These results demonstrate the potential of CBB-augmented, liposome-RNP systems for therapeutic gene editing in the eye, paving the way for single-dose precision medicines to treat IRDs.

Triggering protease-activated cell death is a promising strategy for cancer treatment. Here, we used phage-assisted evolution to reprogram botulinum neurotoxin serotype X proteases to cleave and activate procaspase-1 and gasdermin D, key effectors of inflammatory cell death. We also developed an efficient system to broadly characterize the substrate specificity of wild-type and evolved botulinum neurotoxin serotype X protease variants. Evolved proteases triggered robust cell death across multiple cancer cell lines. The gasdermin D-cleaving protease exclusively induced lytic death, whereas the procaspase-1-cleaving variant initiated both lytic and apoptotic cell death. To enable self-delivery into mammalian cells, we reconstitute evolved proteases with a native BoNT translocation domain, selectively killing cultured cancer cells while sparing non-cancerous cells. Expression of the evolved protease targeting caspase-1 reduced tumor growth in a highly drug-resistant tumor mouse model. These findings establish an evolving protease system to modulate inflammatory cell death and highlight the potential of BoNT proteases as programmable tools for targeted cancer therapy.

Prime editing (PE) can make specific local changes to genomic DNA in living systems but its efficient application currently requires extensive optimization of PE guide RNA (pegRNA) sequences. Here we present OptiPrime, a machine learning model of PE efficiency based on current understanding of PE mechanisms. OptiPrime achieves state-of-the-art accuracy on PE efficiency prediction and enables prediction of nicking guide RNA (PE3) and dual pegRNA (twinPE) outcomes. We validate that OptiPrime has learned the determinants of mammalian mismatch repair (MMR) and is well suited for nominating MMR-evasive silent edits that improve PE efficiency. We demonstrate the use of OptiPrime in a variety of prospective therapeutic contexts in primary human and mouse cells. Lastly, we show that OptiPrime can be used to achieve streamlined and efficient in vivo correction of a pathogenic mutation in the brain of a mouse model of KIF1A-associated neurological disorder. We provide a webserver for OptiPrime ( https://optipri.me/ ) as a community resource.

Transplantation of donor hematopoietic stem and progenitor cells (HSPCs) is a well-established curative treatment for various blood and immune diseases, including severe combined immunodeficiency (SCID). However, it comes with significant toxicities, including graft-versus-host disease (GvHD) and tissue damage resulting from the use of genotoxic chemotherapy-containing conditioning regimens. Autologous transplantation using gene-modified HSPCs eliminates GvHD but currently still relies on genotoxic conditioning. Further, gene modification of HSPCs has commonly utilized integrating viruses, which carry the risk of oncogenesis. The ideal therapy would eliminate the risks associated with current hematopoietic stem cell (HSC) gene-modification and conditioning approaches. Here, we combined base editors (BEs), engineered virus-like particles (eVLPs), and non-genotoxic αCD117 antibody-drug conjugate (ADC) conditioning to explore optimal curative treatment of SCID. We generated a Rag2 SCID mouse model with a single point mutation (pm) and corresponding BE. Rag2pm/pm HSPCs were corrected using SpCas9NG-ABE-eVLPs without off-target effects being detected. Even in settings of low editing, transplantation of BE-corrected HSPCs into αCD117-ADC-conditioned mice led to efficient immune cell production in peripheral blood with normal B cell progenitors in the bone marrow. Combining αCD117-ADC conditioning with transplantation of HSPCs that were base edited using eVLPs successfully reversed the SCID phenotype in mice, showcasing a significant advancement in reducing treatment-related toxicities while enabling disease correction.

Point mutations in the KCNJ13 gene cause autosomal recessive childhood blindness, Leber congenital amaurosis (LCA16), by disrupting Kir7.1 channel function. We describe the etiology of the LCA16 retinopathy phenotype in three patients from two unrelated families harboring a homozygous KCNJ13 missense mutation (c.431T>C, p.Leu144Pro). Our in silico prediction and in vitro validation using a human iPSC-derived retinal pigmented epithelium (RPE) model created via lipid nanoparticle-mediated delivery of the adenine base editor (ABE8e) demonstrated that the L144P mutation impairs Kir7.1 channel function and confirmed that non-viral biologic delivery is clinically translatable. We used two cytosine base editors (CBEs, BE4max-WTCas9 and evoCDA-SpCas9-NG) to correct this mutation in an L144P HEK293 stable cell model, achieving high on-target editing efficiency. However, our electrophysiological measurements showed minimal functional rescue of the channel in CBE-edited cells due to bystander nucleotide editing. Editing with evoCDA introduced a bystander missense mutation (L143F), whereas BE4max primarily generated silent mutations. Extended characterization of BE4max-edited cells revealed a distorted mRNA structure, altered half-life, and reduced abundance of cognate tRNA, all associated with these silent changes. In contrast, prime editing successfully restored channel function. Prime editors targeting the L144P locus achieved approximately 20% on-target editing without introducing bystander nucleotide editing or synonymous changes. Functional assessment demonstrated a strong genotype-phenotype correlation, with restored Kir7.1 channel activity observed in 28% of edited cells (12/43). Overall, these results highlight the importance of comprehensive functional validation of genome editing outcomes and emphasize the need for rigorous preclinical studies to translate therapeutic genome editing into first-in-human trials for genetically diverse diseases.

The performance of prime-editing (PE) systems has been improved by systematic engineering of their protein and small RNA components but the structured RNA motifs appended to the 3' end of PE guide RNAs (pegRNAs)-a key determinant of pegRNA stability and editing efficiency-have not been extensively studied. We introduce PE-PRISM, a high-throughput pooled screen to identify and optimize these 3' RNA motifs in human cells. Here, using PE-PRISM, we evaluated 2,858 RNA motifs across four iterative libraries, including natural and engineered pseudoknots, G-quadruplexes and reverse transcriptase recruitment elements. We applied structure-guided mutagenesis and combinatorial variant screening to refine hits, culminating in the engineered and evolved pseudoknot variants tevo2.0, eHAV and eSBRMV1-A. In a screen correcting 847 pathogenic ClinVar variants, the top-performing motifs improved PE efficiency over the widely used tevopreQ1 motif for >90% of edits. They also increased PE efficiencies for correcting disease-associated mutations in primary human cells and in vivo in mouse brain and liver.

Dravet syndrome (DS) is a severe neurodevelopmental disorder characterized by drug-resistant epilepsy, temperature-sensitive seizures, cognitive impairment, and a high incidence of sudden unexpected death in epilepsy (SUDEP). DS is caused by loss-of-function variants in SCN1A, which encodes the α subunit of the voltage-gated sodium channel (Nav1.1). Current approved treatments manage symptoms of DS but do not correct the root cause of the disease. Here, we describe the use of an adenine base editor (ABE) to directly correct SCN1AR613X, a recurrent variant found in patients with DS. We identified ABE strategies to efficiently correct R613X in engineered homozygous SCN1AR613X human embryonic kidney 293T and mouse Neuro-2a cells (72 and 92% correction efficiencies, respectively). We then used a dual-adeno-associated virus serotype 9 (AAV9) approach to deliver an optimized ABE system to Scn1aR613X/+ mice, which recapitulate several key DS pathologies. AAV9-ABE treatment of Scn1aR613X/+ neonates resulted in efficient DNA and mRNA editing (59 and 97%, respectively, in bulk neocortices), restoring parvalbumin-expressing inhibitory neuron excitability and sodium current to wild-type levels. This ameliorated both spontaneous and temperature-induced seizures and led to a 3.3-fold improvement in 45-day survival over vehicle-treated mice (ABE treated, 90%; and vehicle treated, 27%). Last, ABE treatment in 12-day-old mice resulted in a 3.0-fold improvement in 60-day survival over vehicle-treated mice (ABE treated, 82%; and vehicle treated, 27%). In conclusion, these data validate a strategy to correct SCN1A variants with ABE and highlight the potential of precision genome editing treatments for the treatment of DS and possibly other neurodevelopmental disorders.

Clinical evidence demonstrates that ex vivo gene therapy and genome engineering of hematopoietic stem and progenitor cells (HSPCs) could represent one-time cures. However, while genome editing itself has become increasingly efficient and precise, the toxic conditioning required for hematopoietic stem cell transplantation remains a major barrier to broad clinical implementation of these otherwise curative therapies. In particular, the use of busulfan for myeloablative conditioning constitutes a major safety concern. While preclinical studies established CD117 as a promising target for antigen-specific therapy, clinical translation faced setbacks balancing efficacy and safety. To overcome current limitations, we generated a new CD117-blocking monoclonal antibody (CIM058) and demonstrate its potency to block wild-type HSPCs. To enable long-term blockade of host HSPCs even after transplantation, we used prime editing to engineer CIM058-resistant human CD34+ HSPCs. When combined, CIM058 and the epitope engineered CD34+ HSPCs ameliorated disease phenotype in a β-thalassemia model. Our results suggest that this approach may overcome the reliance on busulfan or other myeloablative conditioning regimens with their associated morbidities, and by enabling toxin-free conditioning and in vivo selection of edited cells, may facilitate clinical implementation of these highly valuable genetic therapies.

The de novo design of small-molecule-binding proteins holds great promise as a potential tool to develop sensors on-demand for arbitrary small molecules. Here we combine deep learning and physics-based methods to generate a family of proteins with diverse and designable pocket geometries, which we employ to computationally design binders for six small-molecule targets. Biophysical characterization of the designed binders reveals nanomolar to low micromolar binding affinities and atomic-level design accuracy. Additionally, we use a cortisol binder to design a chemically induced dimerization (CID) system that enables the construction of a biosensor for cortisol detection. The approach described here demonstrates the potential of the NTF2 fold and deep learning-based protein design in sensor development, paving the way for future platforms to design binders and sensors for small molecules across analytical, environmental, and biomedical applications.

In February 2026, the US Food and Drug Administration (FDA) published a draft guidance on a new plausible mechanism framework for the development and approval of individualized therapies for genetic conditions. Here, we report initial proof-of-concept studies supporting a customizable prime editing platform geared to the treatment of 7 urea cycle disorders (UCDs) and other liver-centered disorders, as well as the outcome of a formal meeting with the FDA to discuss the use of the platform in an “umbrella-of-umbrellas” clinical trial including subjects with any of the 7 UCDs. We anticipate our findings will be of interest to academic investigators and industry sponsors who wish to pursue expeditious FDA approvals of therapies for ultra-rare diseases using the plausible mechanism framework.

Although prime editing (PE) can effect virtually any specified local change to genomic DNA in living systems, its efficient application currently requires extensive optimization of prime editing guide RNA (pegRNA) sequences. We present OptiPrime, a machine learning model of PE efficiency based on our current understanding of the mechanism of prime editing. OptiPrime achieves state-of-the-art accuracy on PE efficiency prediction and also enables prediction of nicking guide RNA (PE3) and dual pegRNA (twinPE) outcomes. We validated that OptiPrime has learned the determinants of mammalian mismatch repair (MMR), and is therefore well suited for nominating MMR-evasive silent edits that improve PE efficiency. We demonstrate the utility of OptiPrime in a variety of prospective therapeutic contexts, including in primary human and mouse cells. Finally, we show how OptiPrime can be used to achieve highly streamlined and efficient in vivo correction of a pathogenic mutation in the brain of a mouse model of KIF1A -associated neurological disorder.

Hutchinson-Gilford progeria syndrome (HGPS) is a premature aging disorder affecting tissues of mesenchymal origin. Most patients harbor a c.1824C>T/p.G608= variant, commonly described as G608G, in exon 11 of LMNA that leads to aberrant splicing and production of the toxic progerin protein. In addition to cardiovascular, dermal, and adipose tissue deterioration, HGPS mouse models also develop progressive bone dysplasia that occurs in patients. Here we characterize the efficacy of in vivo mutation correction with an adenine base editor (ABE) to rescue structural and functional defects in HGPS transgenic murine bone tissue. Treatment of double-copy transgenic osteoblast cultures with a lentiviral-delivered CRISPR-Cas9 ABE achieved nearly 40% gene correction in vitro, resulting in significant reduction of progerin transcripts and protein, in the absence of selective agents. Furthermore, gene correction improved progeroid osteoblasts' capacity to deposit and mineralize extracellular matrix compared to untreated cultures. In vivo, a single intravenous dose of AAV9-delivered ABE corrected the mutation, achieving ~14%, ~22%, ~10% and < 1% correction in bone by six months of age when administered at P3, P14, 1 and 4 months of age, respectively. Partially rescued bone structural and physical parameters were observed in P14-treated mice with concomitant normalization of gene transcriptional programs and intracellular signaling pathways involved in bone remodeling. This work demonstrates in vivo delivery of a locus-specific DNA base editor to bone tissue, delineates the timing of treatment required for maximum efficacy, and suggests that this system might be tailored for application to other monogenic bone disorders.

ABSTRACT:Sickle cell disease (SCD) is a red blood cell disorder caused by a mutation in the β-globin gene, leading to sickle hemoglobin polymerization under low oxygen conditions. Both CRISPR-Cas9 editing and lentiviral transduction have shown promising clinical outcomes, but it remains unclear which approach is superior. Alternatively, new editing tools such as base editing may also be promising and reduce risks of genotoxicity. To compare these approaches, we studied them in an immunocompromised mouse model. We optimized ex vivo conditions in CD34+ hematopoietic stem and progenitor cells (HSPC) and infused edited SCD HSPC into busulfan-conditioned nonirradiated NOD,B6.SCID Il2rγ-/-KitW41/W41 (NBSGW) mice. Ex vivo analysis confirmed successful editing and transduction. At 16 weeks, bone marrow analysis showed similar human CD45+ cell engraftment across all groups (75%-90%). In the competitive transplantation group, there was a lower amount of B-cell lymphoma/leukemia 11A enhancer editing than base editing and lentiviral transduction. A secondary transplantation model yielded similar results. An antisickling assay showed significantly higher red blood cell sickling reduction in the base editing, transduction, and competitive transplantation groups compared to CRISPR-Cas9. In conclusion, although all methods showed therapeutic potential, base editing and lentiviral transduction provided superior outcomes over CRISPR-Cas9-mediated editing in a competitive murine transplantation model.