Efficient expansion and CRISPR-Cas9-mediated gene correction of patient-derived hepatocytes for treatment of inherited liver diseases.

Cell-based ex vivo gene therapy in solid organs, especially the liver, has proven technically challenging. Here, we report a feasible strategy for the clinical application of hepatocyte therapy. We first generated high-quality autologous hepatocytes through the large-scale expansion of patient-derived hepatocytes. Moreover, the proliferating patient-derived hepatocytes, together with the AAV2.7m8 variant identified through screening, enabled CRISPR-Cas9-mediated targeted integration efficiently, achieving functional correction of pathogenic mutations in FAH or OTC. Importantly, these edited hepatocytes repopulated the injured mouse liver at high repopulation levels and underwent maturation, successfully treating mice with tyrosinemia following transplantation. Our study combines ex vivo large-scale cell expansion and gene editing in patient-derived transplantable hepatocytes, which holds potential for treating human liver diseases.

Fork coupling directs DNA replication elongation and termination.

DNA replication is initiated at multiple loci to ensure timely duplication of eukaryotic genomes. Sister replication forks progress bidirectionally, and replication terminates when two convergent forks encounter one another. To investigate the coordination of replication forks, we developed a replication-associated in situ HiC method to capture chromatin interactions involving nascent DNA. We identify more than 2000 fountain-like structures of chromatin contacts in human and mouse genomes, indicative of coupling of DNA replication forks. Replication fork interaction not only occurs between sister forks but also involves forks from two distinct origins to predetermine replication termination. Termination-associated chromatin fountains are sensitive to replication stress and lead to coupled forks-associated genomic deletions in cancers. These findings reveal the spatial organization of DNA replication forks within the chromatin context.

Cohesin maintains replication timing to suppress DNA damage on cancer genes.

Cohesin loss-of-function mutations are frequently observed in tumors, but the mechanism underlying its role in tumorigenesis is unclear. Here, we found that depletion of RAD21, a core subunit of cohesin, leads to massive genome-wide DNA breaks and 147 translocation hotspot genes, co-mutated with cohesin in multiple cancers. Increased DNA damages are independent of RAD21-loss-induced transcription alteration and loop anchor disruption. However, damage-induced chromosomal translocations coincide with the asymmetrically distributed Okazaki fragments of DNA replication, suggesting that RAD21 depletion causes replication stresses evidenced by the slower replication speed and increased stalled forks. Mechanistically, approximately 30% of the human genome exhibits an earlier replication timing after RAD21 depletion, caused by the early initiation of >900 extra dormant origins. Correspondingly, most translocation hotspot genes lie in timing-altered regions. Therefore, we conclude that cohesin dysfunction causes replication stresses induced by excessive DNA replication initiation, resulting in gross DNA damages that may promote tumorigenesis.

Human 8-cell embryos enable efficient induction of disease-preventive mutations without off-target effect by cytosine base editor.

Approximately 140 million people worldwide are homozygous carriers of APOE4 (ε4), a strong genetic risk factor for late onset familial and sporadic Alzheimer's disease (AD), 91% of whom will develop AD at earlier age than heterozygous carriers and noncarriers. Susceptibility to AD could be reduced by targeted editing of APOE4, but a technical basis for controlling the off-target effects of base editors is necessary to develop low-risk personalized gene therapies. Here, we first screened eight cytosine base editor variants at four injection stages (from 1- to 8-cell stage), and found that FNLS-YE1 variant in 8-cell embryos achieved the comparable base conversion rate (up to 100%) with the lowest bystander effects. In particular, 80% of AD-susceptible ε4 allele copies were converted to the AD-neutral ε3 allele in human ε4-carrying embryos. Stringent control measures combined with targeted deep sequencing, whole genome sequencing, and RNA sequencing showed no DNA or RNA off-target events in FNLS-YE1-treated human embryos or their derived stem cells. Furthermore, base editing with FNLS-YE1 showed no effects on embryo development to the blastocyst stage. Finally, we also demonstrated FNLS-YE1 could introduce known protective variants in human embryos to potentially reduce human susceptivity to systemic lupus erythematosus and familial hypercholesterolemia. Our study therefore suggests that base editing with FNLS-YE1 can efficiently and safely introduce known preventive variants in 8-cell human embryos, a potential approach for reducing human susceptibility to AD or other genetic diseases.

Engineering of efficiency-enhanced Cas9 and base editors with improved gene therapy efficacies.

Editing efficiency is pivotal for the efficacies of CRISPR-based gene therapies. We found that fusing an HMG-D domain to the N terminus of SpCas9 (named efficiency-enhanced Cas9 [eeCas9]) significantly increased editing efficiency by 1.4-fold on average. The HMG-D domain also enhanced the activities of non-NGG PAM Cas9 variants, high-fidelity Cas9 variants, smaller Cas9 orthologs, Cas9-based epigenetic regulators, and base editors in cell lines. Furthermore, we discovered that eeCas9 exhibits comparable off-targeting effects with Cas9, and its specificity could be increased through ribonucleoprotein delivery or using hairpin single-guide RNAs and high-fidelity Cas9s. The entire eeCas9 could be packaged into an adeno-associated virus vector and exhibited a 1.7- to 2.6-fold increase in editing efficiency targeting the Pcsk9 gene in mice, leading to a greater reduction of serum cholesterol levels. Moreover, the efficiency of eeA3A-BE3 also surpasses that of A3A-BE3 in targeting the promoter region of γ-globin genes or BCL11A enhancer in human hematopoietic stem cells to reactivate γ-globin expression for the treatment of ß-hemoglobinopathy. Together, eeCas9 and its derivatives are promising editing tools that exhibit higher activity and therapeutic efficacy for both in vivo and ex vivo therapeutics.

Safeguarding genome integrity during gene-editing therapy in a mouse model of age-related macular degeneration.

Ensuring genome safety during gene editing is crucial for clinical translation of the high-efficient CRISPR-Cas9 toolbox. Therefore, the undesired events including chromosomal translocations, vector integrations, and large deletions arising during therapeutic gene editing remain to be adequately addressed or tackled in vivo. Here, we apply CRISPR-Cas9TX in comparison to CRISPR-Cas9 to target Vegfa for the treatment of age-related macular degeneration (AMD) disease in a mouse model. AAV delivery of both CRISPR-Cas9 and CRISPR-Cas9TX can efficiently inhibit laser-induced neovascularization. Importantly, Cas9TX almost eliminates chromosomal translocations that occur at a frequency of approximately 1% in Cas9-edited mouse retinal cells. Strikingly, the widely observed AAV integration at the target Vegfa site is also greatly reduced from nearly 50% of edited events to the background level during Cas9TX editing. Our findings reveal that chromosomal structural variations routinely occur during in vivo genome editing and highlight Cas9TX as a superior form of Cas9 for in vivo gene disruption.

CRISPR/Cas9-induced structural variations expand in T lymphocytes in vivo.

CRISPR/Cas9 has been adapted to disrupt endogenous genes in adoptive T-lymphocyte therapy to prevent graft-versus-host disease. However, genome editing also generates prevalent deleterious structural variations (SVs), including chromosomal translocations and large deletions, raising safety concerns about reinfused T cells. Here, we dynamically monitored the progression of SVs in a mouse model of T-cell receptor (TCR)-transgenic T-cell adoptive transfer, mimicking TCR T therapeutics. Remarkably, CRISPR/Cas9-induced SVs persist and undergo clonal expansion in vivo after three weeks or even two months, evidenced by high enrichment and low junctional diversity of identified SVs post infusion. Specifically, we detected 128 expanded translocations, with 20 615 as the highest number of amplicons. The identified SVs are stochastically selected among different individuals and show an inconspicuous locus preference. Similar to SVs, viral DNA integrations are routinely detected in edited T cells and also undergo clonal expansion. The persistent SVs and viral DNA integrations in the infused T cells may constantly threaten genome integrity, drawing immediate attention to the safety of CRISPR/Cas9-engineered T cells mediated immunotherapy.

Comprehensive assessment of miniature CRISPR-Cas12f nucleases for gene disruption.

Because of their small size, the recently developed CRISPR-Cas12f nucleases can be effectively packaged into adeno-associated viruses for gene therapy. However, a systematic evaluation of the editing outcomes of CRISPR-Cas12f is lacking. In this study, we apply a high-throughput sequencing method to comprehensively assess the editing efficiency, specificity, and safety of four Cas12f proteins in parallel with that of Cas9 and two Cas12a proteins at multiple genomic sites. Cas12f nucleases achieve robust cleavage at most of the tested sites and mainly produce deletional fragments. In contrast, Cas9 and Cas12a show relatively higher editing efficiency at the vast majority of the tested sites. However, the off-target hotspots identified in the Cas9- and Cas12a-edited cells are negligibly detected in the Cas12f-edited cells. Moreover, compared to Cas9 and Cas12a nucleases, Cas12f nucleases reduce the levels of chromosomal translocations, large deletions, and integrated vectors by 2- to 3-fold. Therefore, our findings confirm the editing capacity of Cas12f and reveal the ability of this nuclease family to preserve genome integrity during genome editing.

CTCF DNA-binding domain undergoes dynamic and selective protein-protein interactions.

CTCF is a predominant insulator protein required for three-dimensional chromatin organization. However, the roles of its insulation of enhancers in a 3D nuclear organization have not been fully explained. Here, we found that the CTCF DNA-binding domain (DBD) forms dynamic self-interacting clusters. Strikingly, CTCF DBD clusters were found to incorporate other insulator proteins but are not coenriched with transcriptional activators in the nucleus. This property is not observed in other domains of CTCF or the DBDs of other transcription factors. Moreover, endogenous CTCF shows a phenotype consistent with the DBD by forming small protein clusters and interacting with CTCF motif arrays that have fewer transcriptional activators bound. Our results reveal an interesting phenomenon in which CTCF DBD interacts with insulator proteins and selectively localizes to nuclear positions with lower concentrations of transcriptional activators, providing insights into the insulation function of CTCF.

The origin of unwanted editing byproducts in gene editing.

The rapid development of CRISPR-Cas genome editing tools has greatly changed the way to conduct research and holds tremendous promise for clinical applications. During genome editing, CRISPR-Cas enzymes induce DNA breaks at the target sites and subsequently the DNA repair pathways are recruited to generate diverse editing outcomes. Besides off-target cleavage, unwanted editing outcomes including chromosomal structural variations and exogenous DNA integrations have recently raised concerns for clinical safety. To eliminate these unwanted editing byproducts, we need to explore the underlying mechanisms for the formation of diverse editing outcomes from the perspective of DNA repair. Here, we describe the involved DNA repair pathways in sealing Cas enzyme-induced DNA double-stranded breaks and discuss the origins and effects of unwanted editing byproducts on genome stability. Furthermore, we propose the potential risk of inhibiting DNA repair pathways to enhance gene editing. The recent combined studies of DNA repair and CRISPR-Cas editing provide a framework for further optimizing genome editing to enhance editing safety.

Long-term correction of hemophilia B through CRISPR/Cas9 induced homology-independent targeted integration.

CRISPR/Cas9-mediated site-specific insertion of exogenous genes holds potential for clinical applications. However, it is still infeasible because homologous recombination (HR) is inefficient, especially for non-dividing cells. To overcome the challenge, we report that a homology-independent targeted integration (HITI) strategy is used for permanent integration of high-specificity-activity Factor IX variant (F9 Padua, R338L) at the albumin (Alb) locus in a novel hemophilia B (HB) rat model. The knock-in efficiency reaches 3.66%, as determined by droplet digital PCR (ddPCR). The clotting time is reduced to a normal level four weeks after treatment, and the circulating factor IX (FIX) level is gradually increased up to 52% of the normal level over nine months even after partial hepatectomy, demonstrating the amelioration of hemophilia. Through primer-extension-mediated sequencing (PEM-seq), no significant off-target effect is detected. This study not only provides a novel model for HB but also identifies a promising therapeutic approach for rare inherited diseases.

PEM-seq comprehensively quantifies DNA repair outcomes during gene-editing and DSB repair.

The repair products of double-stranded DNA breaks (DSBs) are crucial for investigating the mechanism underlying DNA damage repair as well as evaluating the safety and efficiency of gene-editing; however, a comprehensively quantitative assay remains to be established. Here, we describe the step-by-step instructions of the primer extension-mediated sequencing (PEM-seq), followed by the framework of data processing and statistical analysis. PEM-seq presents a full spectrum of repair outcomes for both genome-editing-induced and endogenous DSBs in mouse and human cells. For complete details on the use and execution of this profile, please refer to Gan et al. (2021), Yin et al. (2019), Liu et al. (2021a), and Zhang et al. (2021).

C-terminal deletion-induced condensation sequesters AID from IgH targets in immunodeficiency.

In activated B cells, activation-induced cytidine deaminase (AID) generates programmed DNA lesions required for antibody class switch recombination (CSR), which may also threaten genome integrity. AID dynamically shuttles between cytoplasm and nucleus, and the majority stays in the cytoplasm due to active nuclear export mediated by its C-terminal peptide. In immunodeficient-patient cells expressing mutant AID lacking its C-terminus, a catalytically active AID-delC protein accumulates in the nucleus but nevertheless fails to support CSR. To resolve this apparent paradox, we dissected the function of AID-delC proteins in the CSR process and found that they cannot efficiently target antibody genes. We demonstrate that AID-delC proteins form condensates both in vivo and in vitro, dependent on its N-terminus and on a surface arginine-rich patch. Co-expression of AID-delC and wild-type AID leads to an unbalanced nuclear AID-delC/AID ratio, with AID-delC proteins able to trap wild-type AID in condensates, resulting in a dominant-negative phenotype that could contribute to immunodeficiency. The co-condensation model of mutant and wild-type proteins could be an alternative explanation for the dominant-negative effect in genetic disorders.

Cas9 exo-endonuclease eliminates chromosomal translocations during genome editing.

The mechanism underlying unwanted structural variations induced by CRISPR-Cas9 remains poorly understood, and no effective strategy is available to inhibit the generation of these byproducts. Here we find that the generation of a high level of translocations is dependent on repeated cleavage at the Cas9-targeting sites. Therefore, we employ a strategy in which Cas9 is fused with optimized TREX2 to generate Cas9TX, a Cas9 exo-endonuclease, which prevents perfect DNA repair and thereby avoids repeated cleavage. In comparison with CRISPR-Cas9, CRISPR-Cas9TX greatly suppressed translocation levels and enhanced the editing efficiency of single-site editing. The number of large deletions associated with Cas9TX was also reduced to very low level. The application of CRISPR-Cas9TX for multiplex gene editing in chimeric antigen receptor T cells nearly eliminated deleterious chromosomal translocations. We report the mechanism underlying translocations induced by Cas9, and propose a general strategy for reducing chromosomal abnormalities induced by CRISPR-RNA-guided endonucleases.

Improving the efficiency of CRISPR-Cas12a-based genome editing with site-specific covalent Cas12a-crRNA conjugates.

The CRISPR-Cas12a system shows unique features compared with widely used Cas9, making it an attractive and potentially more precise alternative. However, the adoption of this system has been hindered by its relatively low editing efficiency. Guided by physical chemical principles, we covalently conjugated 5' terminal modified CRISPR RNA (crRNA) to a site-specifically modified Cas12a through biorthogonal chemical reaction. The genome editing efficiency of the resulting conjugated Cas12a complex (cCas12a) was substantially higher than that of the wild-type complex. We also demonstrated that cCas12a could be used for precise gene knockin and multiplex gene editing in a chimeric antigen receptor T cell preparation with efficiency much higher than that of the wild-type system. Overall, our findings indicate that covalently linking Cas nuclease and crRNA is an effective approach to improve the Cas12a-based genome editing system and could potentially provide an insight into engineering other Cas family members with low efficiency as well.

RAG2 abolishes RAG1 aggregation to facilitate V(D)J recombination.

RAG1 and RAG2 form a tetramer nuclease to initiate V(D)J recombination in developing T and B lymphocytes. The RAG1 protein evolves from a transposon ancestor and possesses nuclease activity that requires interaction with RAG2. Here, we show that the human RAG1 aggregates in the nucleus in the absence of RAG2, exhibiting an extremely low V(D)J recombination activity. In contrast, RAG2 does not aggregate by itself, but it interacts with RAG1 to disrupt RAG1 aggregates and thereby activate robust V(D)J recombination. Moreover, RAG2 from mouse and zebrafish could not disrupt the aggregation of human RAG1 as efficiently as human RAG2 did, indicating a species-specific regulatory mechanism for RAG1 by RAG2. Therefore, we propose that RAG2 coevolves with RAG1 to release inert RAG1 from aggregates and thereby activate V(D)J recombination to generate diverse antigen receptors in lymphocytes.

Global detection of DNA repair outcomes induced by CRISPR-Cas9.

CRISPR-Cas9 generates double-stranded DNA breaks (DSBs) to activate cellular DNA repair pathways for genome editing. The repair of DSBs leads to small insertions or deletions (indels) and other complex byproducts, including large deletions and chromosomal translocations. Indels are well understood to disrupt target genes, while the other deleterious byproducts remain elusive. We developed a new in silico analysis pipeline for the previously described primer-extension-mediated sequencing assay to comprehensively characterize CRISPR-Cas9-induced DSB repair outcomes in human or mouse cells. We identified tremendous deleterious DSB repair byproducts of CRISPR-Cas9 editing, including large deletions, vector integrations, and chromosomal translocations. We further elucidated the important roles of microhomology, chromosomal interaction, recurrent DSBs, and DSB repair pathways in the generation of these byproducts. Our findings provide an extra dimension for genome editing safety besides off-targets. And caution should be exercised to avoid not only off-target damages but also deleterious DSB repair byproducts during genome editing.

Transcription shapes DNA replication initiation to preserve genome integrity.

BACKGROUND: Early DNA replication occurs within actively transcribed chromatin compartments in mammalian cells, raising the immediate question of how early DNA replication coordinates with transcription to avoid collisions and DNA damage. RESULTS: We develop a high-throughput nucleoside analog incorporation sequencing assay and identify thousands of early replication initiation zones in both mouse and human cells. The identified early replication initiation zones fall in open chromatin compartments and are mutually exclusive with transcription elongation. Of note, early replication initiation zones are mainly located in non-transcribed regions adjacent to transcribed regions. Mechanistically, we find that RNA polymerase II actively redistributes the chromatin-bound mini-chromosome maintenance complex (MCM), but not the origin recognition complex (ORC), to actively restrict early DNA replication initiation outside of transcribed regions. In support of this finding, we detect apparent MCM accumulation and DNA replication initiation in transcribed regions due to anchoring of nuclease-dead Cas9 at transcribed genes, which stalls RNA polymerase II. Finally, we find that the orchestration of early DNA replication initiation by transcription efficiently prevents gross DNA damage. CONCLUSION: RNA polymerase II redistributes MCM complexes, but not the ORC, to prevent early DNA replication from initiating within transcribed regions. This RNA polymerase II-driven MCM redistribution spatially separates transcription and early DNA replication events and avoids the transcription-replication initiation collision, thereby providing a critical regulatory mechanism to preserve genome stability.

In-depth assessment of the PAM compatibility and editing activities of Cas9 variants.

A series of Cas9 variants have been developed to improve the editing fidelity or targeting range of CRISPR-Cas9. Here, we employ a high-throughput sequencing approach primer-extension-mediated sequencing to analyze the editing efficiency, specificity and protospacer adjacent motif (PAM) compatibility of a dozen of SpCas9 variants at multiple target sites in depth, and our findings validate the high fidelity or broad editing range of these SpCas9 variants. With regard to the PAM-flexible SpCas9 variants, we detect significantly increased levels of off-target activity and propose a trade-off between targeting range and editing specificity for them, especially for the near-PAM-less SpRY. Moreover, we use a deep learning model to verify the consistency and predictability of SpRY off-target sites. Furthermore, we combine high-fidelity SpCas9 variants with SpRY to generate three new SpCas9 variants with both high fidelity and broad editing range. Finally, we also find that the existing SpCas9 variants are not effective in suppressing genome instability elicited by CRISPR-Cas9 editing, raising an urgent issue to be addressed.