Glioblastoma (GBM) is an aggressive brain tumor with limited treatment options and poor patient survival, underscoring the need for novel, to our knowledge, therapeutic strategies and improved preclinical models. Patient-derived tumor spheroids (PDTSs) offer a physiologically relevant in vitro platform for evaluating treatments such as chimeric antigen receptor (CAR) T cell therapy, chemotherapy, and radiation. However, significant challenges remain in monitoring the complex three-dimensional (3D) microenvironment of the GBM PDTSs. Current imaging techniques used for this purpose are primarily endpoint analyses which lack critical real-time, non-invasive capabilities that ultimately preclude longitudinal and continuous monitoring. In this study, we introduce quantitative oblique back-illumination microscopy (qOBM) as a label-free and non-invasive imaging approach for longitudinal and continuous, high-resolution monitoring of GBM PDTSs during treatment. qOBM enables real-time visualization of cellular processes, including apoptosis, cell migration, and T cell-mediated cytotoxicity by leveraging tomographic refractive index-based quantitative imaging. We construct a compact qOBM system that fits within common incubators and apply it to study the effects of radiation, chemotherapy, and immunotherapy on three patient-derived GBM cell lines, extracting both static and dynamic image features over a 72 h treatment period. Additionally, we develop machine learning models to predict spheroid viability and cytotoxicity, demonstrating the potential of qOBM to enhance treatment evaluation. Our findings establish qOBM as a powerful tool for longitudinal and continuous spheroid monitoring, offering a non-destructive, high-resolution alternative to conventional endpoint assays and improving the evaluation of preclinical treatments for GBM.

Human T cells modified with nucleic acids constitute a powerful and emerging therapeutic modality for cancer, autoimmune disorders, and aging related diseases. However, delivering nucleic acids, such as mRNA encoding synthetic receptors, transcription factors, cytokines, or genome editors to T cells can be challenging, as nucleic acids can unintentionally reduce viable T cell yield and function, particularly when delivered sequentially ex vivo. To address this challenge, we evaluated the efficiency of serial non-viral delivery of synthetic mRNA encapsulated within lipid nanoparticles (LNPs) incorporating cationic lipid, 1,2-Dioleoyl-3-trimethylammonium-propane (DOTAP), and fusogenic helper lipid, 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE). DOTAP/DOPE-containing LNPs delivered mRNA more efficiently than a clinically benchmarked 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC)-containing LNP formulation, resulting in significantly higher protein expression in primary human T cells. Furthermore, DOTAP/DOPE-LNP delivery resulted in 100% higher yield of live cells compared to electroporation, an advantage that compounded over serial rounds, with LNP-treated cultures maintaining substantially more viable cells through all four transfections. Additionally, serial transfection of mRNAs encoding a therapeutically relevant chimeric antigen receptor (CAR) payload produced functional CAR T cells. These results demonstrate that LNPs can be a viable platform for serial and iterative T cell engineering using multiple mRNA payloads, providing an alternative to electroporation for the engineering of therapeutic T cells.

To systematically map cellular factors constraining nonviral genome editing, influencing uptake and intracellular trafficking, we develop a genome-wide CRISPR screening platform linking perturbation of 19,114 genes to editing outcomes in human cells. We identify six negative regulators of delivery whose depletion increases editing efficiency by up to six-fold across diverse payloads, loci, and cell types. We test the top two factors, GJB2 and BET1L, in two distinct human models: correction of a pathogenic adenine base mutation in KCNJ13 and introduction of a cytosine base mutation in the GABAA receptor gene. Depletion of either improves base-editing outcomes by 6-fold, potentially through effects on delivery. In a patient-derived model of retinal channelopathy, knockdown of either gene improves lipid nanoparticle base editing efficiency by over 3.5-fold. This enables functional restoration of Kir7.1 ion channels in a subset of edited cells, highlighting cellular barriers as actionable targets to enhance the potency of genetic therapies.

The urokinase-type plasminogen activator receptor (uPAR) is a GPI-anchored cell surface protein that regulates leukocyte adhesion, migration, and activation, thereby contributing to inflammation and tissue remodeling. However, its role in Alzheimer's disease (AD), particularly in relation to glial dysfunction, remains poorly defined. Here, we investigated the temporal and spatial regulation of uPAR expression across AD mouse models with intact or deficient adaptive immunity. Using immunohistochemistry, we assessed uPAR expression in Rag2/Il2rg-/- (Rag), Rag2/Il2rg-/--5xFAD (Rag-5xFAD), C57BL/6 (WT), and 5xFAD mice across multiple brain regions. uPAR expression increased with age and was significantly elevated in 5xFAD mice, with robust upregulation evident by 6 months irrespective of immune status. Immunofluorescence revealed that uPAR localized predominantly to Iba1+ microglia clustered around Aβ plaques, with limited neuronal expression. Bulk RNA sequencing of Rag-5xFAD brain tissue demonstrated enrichment of disease-associated microglia (DAM) and senescence-related transcriptional programs. These findings indicate that uPAR marks a subset of plaque-associated glial cells undergoing functional and transcriptional remodeling in AD, independent of peripheral adaptive immune signaling. Collectively, our results identify uPAR as a marker of dysfunctional, DAM-like microglia and implicate it in senescence-associated neuroinflammatory pathways. This work provides a framework for future studies targeting uPAR-expressing glial populations as a potential therapeutic strategy in AD.

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.

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.

Biofabrication and biomedical manufacturing are inherently multidisciplinary, integrating living systems with advanced manufacturing to create functional products for applications spanning regenerative engineering and medicine, in vitro disease modeling, drug discovery, and medical devices. As these technologies develop, they are emerging as core enablers of next-generation healthcare and life-science innovation. In the United States (U.S.), rapid progress across fabrication processes, material systems, physics-based modeling, and translation-oriented strategies is expanding the achievable design space and accelerating movement from laboratory demonstrations toward clinical and commercial deployment. We introduce major U.S. research frontiers and highlight representative advances in this field that support applications including organoids and other microphysiological systems for in vitro testing, engineered tissue constructs for in vivo use, and medical devices and biohybrid platforms. We further provide an outlook on advancing robust, ethical biofabrication and biomedical manufacturing in the U.S. research ecosystem.

CRISPR-Cas9-based genome editors can precisely target and edit genes efficiently. However, prolonged Cas9 activity poses challenges for laboratory experiments and raises safety concerns for therapeutic applications due to unintended consequences such as off-target editing, genotoxicity, immunogenicity, and undesired on-target modifications. Here, we evaluate a novel molecular glue degradation system, called Cas9-degron (Cas9-d), designed to degrade Cas9 in the presence of the US Food and Drug Administration (FDA)-approved drug, pomalidomide (POM). This system is highly biocompatible and rapidly reduces Cas9 protein levels within 4 h of induction, resulting in a 3- to 5-fold decrease in editing at on-target sites. The reduction is reversible, as Cas9 levels are restored within 24 h after POM withdrawal. Without initiating degradation, the on-target editing efficiency and accuracy of the Cas9-d system remain intact in different human cell types, including hepatic cell lines and human induced pluripotent stem cell (hiPSC)-derived GABAergic neurons. Cells edited with the Cas9-d system were healthy and functional, exhibiting minimal toxicity from using the strategy. The Cas9-d system provides a versatile approach to adjust Cas9 levels, demonstrating its potential as an experimental tool for controlling genome editing outcomes in vitro and ex vivo. With further development, it holds promise for enhancing somatic cell genome editing in vivo.

Gene-editing technologies promise to create a new class of therapeutics that can achieve permanent correction with a single intervention. Besides eliminating mutant alleles in familial disease, gene-editing can also be used to favorably manipulate upstream pathophysiologic events and alter disease-course in wider patient populations, but few such feasible therapeutic avenues have been reported. Here we use CRISPR-Cas9 to edit the last exon of amyloid precursor protein (App), relevant for Alzheimer's disease (AD). Our strategy effectively eliminates an endocytic (YENPTY) motif at APP C-terminus, while preserving the N-terminus and compensatory APP-homologues. This manipulation favorably alters events along the amyloid-pathway - inhibiting toxic APP-β-cleavage fragments (including Aβ) and upregulating neuroprotective APP-α-cleavage products. AAV-driven editing ameliorates neuropathologic, electrophysiologic, and behavioral deficits in an AD knockin mouse model. Effects persist for many months, and no abnormalities are seen in WT mice even after germline App-editing; underlining overall efficacy and safety. Pathologic alterations in the glial-transcriptome of App-KI mice, as seen by single nuclei RNA-sequencing (sNuc-Seq), are also normalized by App C-terminus editing. Our strategy takes advantage of innate transcriptional rules that render terminal exons insensitive to nonsense-decay, and the upstream manipulation is expected to be effective for all forms of AD. These studies offer a path for a one-time disease-modifying treatment for AD.

Genetic medicines, including CRISPR/Cas technologies, extend tremendous promise for addressing unmet medical need in inherited retinal disorders and other indications; however, there remain challenges for the development of therapeutics. Herein, we evaluate genome editing by engineered Cas9 ribonucleoproteins (eRNP) in vivo via subretinal administration using mouse and pig animal models. Subretinal administration of adenine base editor and double strand break-inducing Cas9 nuclease eRNPs mediate genome editing in both species. Editing occurs in retinal pigmented epithelium (RPE) and photoreceptor cells, with favorable tolerability in both species. Using transgenic reporter strains, we determine that editing primarily occurs close to the site of administration, within the bleb region associated with subretinal injection. Our results show that subretinal administration of BE-eRNPs in mice mediates base editing of up to 12% of the total neural retina, with an average rate of 7% observed at the highest dose tested. In contrast, a substantially lower editing efficiency was observed in minipigs; even with direct quantification of only the treated region, a maximum base editing rate of 1.5%, with an average rate of <1%, was observed. Our data highlight the importance of species consideration in preclinical studies for the development of genetic medicines targeting the eye and provide an example of a lack of translation between small and larger animal models in the context of subretinal administration of Cas9 eRNPs.

Chimeric antigen receptor (CAR) T cell therapy for solid tumours is challenging because of the immunosuppressive tumour microenvironment and a complex manufacturing process. Cellular manufacturing protocols directly impact CAR T cell yield, phenotype and metabolism, which correlates with in vivo potency and persistence. Although metabolic fitness is a critical quality attribute, how T cell metabolic requirements vary throughout the manufacturing process remains unexplored. Here we use optical metabolic imaging (OMI), a non-invasive, label-free method to evaluate single-cell metabolism. Using OMI, we identified the impacts of media composition on CAR T cell metabolism, activation strength and kinetics, and phenotype. We demonstrate that OMI parameters can indicate cell cycle stage and optimal gene transfer conditions for both viral transduction and electroporation-based CRISPR/Cas9. In a CRISPR-edited anti-GD2 CAR T cell model, OMI measurements allow accurate prediction of an oxidative metabolic phenotype that yields higher in vivo potency against neuroblastoma. Our data support OMI as a robust, sensitive analytical tool to optimize manufacturing conditions and monitor cell metabolism for increased CAR T cell yield and metabolic fitness. Monitoring T cell metabolism during chimeric antigen receptor T manufacturing using optical metabolic imaging enables fine-tuning of manufacturing conditions to increase T cell yield and fitness.

Chimeric antigen receptor (CAR) T cells have limited efficacy against solid tumors including neuroblastoma. Here, we evaluated whether low-dose radiation delivered by radiopharmaceutical therapy (RPT), known to potentiate immune checkpoint inhibitors, can synergize with CRISPR-edited GD2 TRAC-CAR T cells to improve outcomes in neuroblastoma. We found that in the localized model of neuroblastoma, low-dose radiation delivered by 177Lu-NM600, an alkylphosphocholine mimetic RPT agent, followed 9 days later by GD2 TRAC-CAR T cells led to complete tumor regression. Irradiation of neuroblastoma before GD2 TRAC-CAR T cells enhanced the release by CAR T cells of perforin, granzyme B, tumor necrosis factor-α, and interleukin-7 while abrogating transforming growth factor-β1. Low-dose RPT up-regulated the death receptor Fas on neuroblastoma, potentially enabling CAR-independent killing. This suggests that low-dose RPT can enhance suboptimal CAR T cell efficacy against solid tumors. However, optimization of radiation dose and timing may be needed for each patient and RPT agent to account for varied tumor radiosensitivity and dosimetry.

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.

This article complements the 2021 International Society for Stem Cell Research (ISSCR) Guidelines for Stem Cell Research and Clinical Translation by explaining what "adequate and appropriate scientific justification" means for human pluripotent stem cell, embryo, and related research, thus providing practical advice for review and oversight bodies, regulators, and investigators who conduct research in these areas.

How should we govern our increasing power to intervene in the processes of life? Genome editing, especially of the human germline, has brought this question to the forefront of global debate. We must seek to rectify shortcomings of earlier deliberative approaches by setting aside a science-and-technology first approach; expanding the range of questions for deliberation; revisiting the distribution of innovation's benefits and risks; and reimagining the limits of research. This Perspective from the Organizing Committee of the 2025 Global Observatory for Genome Editing International Summit calls for a new social compact, recognizing and rendering accountable the constitutive role of science and technology in shaping the meaning of human life in the 21st century.

After administering genome editors, their efficiency is limited by a multi-step process involving cellular uptake, trafficking, and nuclear import of the vector and its payload. These processes vary widely across cell types and differ depending on the nature and structure of the vector, whether it is a lipid nanoparticle or a different synthetic material. We developed a novel genome-wide CRISPR screening strategy to better understand these limitations within human cells to identify genes modulating cellular uptake, payload delivery, and gene editing efficiency. Our screen interrogates the cellular processes controlling genome editing by Cas-based nuclease and base editing strategies in human cells. We designed a genome-wide screen targeting 19,114 genes in HEK293 cells, and we identified six genes whose knockout increased nonviral editing efficiency in human cells by up to five-fold. Further validation through arrayed knockouts of the top hits from our screen boosted the editing efficiency from 5% to 50% when Cas9 was delivered via lipid-based nanoparticles. By designing the guides to target the screen library cassette, we could accurately track the library sgRNA identity and the editing outcome on the same amplicon via short-read sequencing, enabling the identification of rare outcomes via 'computationally' sorting edited from unedited cells within a heterogenous pool of >200M cells. In patient-derived human retinal pigment epithelium cells derived from pluripotent stem cells, BET1L, GJB2, and MS4A13 gene knockouts increased targeted genome editing by over five-fold. We anticipate that this high-throughput screening approach will facilitate the systematic engineering of novel nonviral genome editing delivery methods, where the identified novel gene hits can be further used to increase editing efficiency for other therapeutically relevant cell types.

The KCNJ13 gene encodes the Kir7.1 protein, and mutations cause Leber’s Congenital Amaurosis (LCA) and Snowflake Vitreoretinal Degeneration (SVD), leading to early-onset vision loss. One such nonsense mutation, R166X, results in nystagmus, poor night vision, and visual impairment. We examined therapeutic options for the R166X nonsense point mutation. Unlike missense genetic mutations, which can be rectified by gene augmentation and genome editing, nonsense mutations provide an opportunity to test additional readthrough therapies. Attempts to produce human induced pluripotent stem cells (hiPSCs) with the R166X mutation (CGA to TGA) were unsuccessful. This may be due to challenges in chromatin structure and folding that hinder access to the targeted loci. Therefore, we constructed an open reading frame (ORF) stably integrated HEK293T line by inserting the wild type (WT) or R166X-KCNJ13 gene using the FLP-FRT recombinase technique. While genome editing strategies did not repair the R166X mutation efficiently in this cell line, an arginine anticodon-engineered tRNA (ACE-tRNAArg.UGA) restored K+ channel expression and function. This study provides an example in precision medicine where translational readthrough strategies can rescue channel function at a mutation that is difficult to correct via genome editing.