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CRISPR/Cas9-mediated genome engineering: an adeno-associated viral (AAV) vector toolbox.
Its remarkable ease and efficiency make the CRISPR (clustered regularly interspaced short palindromic repeats) DNA editing machinery highly attractive as a new tool for experimental gene annotation and therapeutic genome engineering in eukaryotes. Here, we report a versatile set of plasmids and vectors derived from adeno-associated virus (AAV) that allow robust and specific delivery of the two essential CRISPR components - Cas9 and chimeric g(uide)RNA - either alone or in combination. All our constructs share a modular design that enables simple and stringent guide RNA (gRNA) cloning as well as rapid exchange of promoters driving Cas9 or gRNA. Packaging into potent synthetic AAV capsids permits CRISPR delivery even into hard-to-transfect targets, as shown for human T-cells. Moreover, we demonstrate the feasibility to direct Cas9 expression to or away from hepatocytes, using a liver-specific promoter or a hepatic miRNA binding site, respectively. We also report a streamlined and economical protocol for detection of CRISPR-induced mutations in less than 3 h. Finally, we provide original evidence that AAV/CRISPR vectors can be exploited for gene engineering in vivo, as exemplified in the liver of adult mice. Our new tools and protocols should foster the broad application of CRISPR technology in eukaryotic cells and organisms, and accelerate its clinical translation into humans.
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Base-edited CAR T cells for combinational therapy against T cell malignancies.
Targeting T cell malignancies using chimeric antigen receptor (CAR) T cells is hindered by 'T v T' fratricide against shared antigens such as CD3 and CD7. Base editing offers the possibility of seamless disruption of gene expression of problematic antigens through creation of stop codons or elimination of splice sites. We describe the generation of fratricide-resistant T cells by orderly removal of TCR/CD3 and CD7 ahead of lentiviral-mediated expression of CARs specific for CD3 or CD7. Molecular interrogation of base-edited cells confirmed elimination of chromosomal translocations detected in conventional Cas9 treated cells. Interestingly, 3CAR/7CAR co-culture resulted in 'self-enrichment' yielding populations 99.6% TCR-/CD3-/CD7-. 3CAR or 7CAR cells were able to exert specific cytotoxicity against leukaemia lines with defined CD3 and/or CD7 expression as well as primary T-ALL cells. Co-cultured 3CAR/7CAR cells exhibited highest cytotoxicity against CD3 + CD7 + T-ALL targets in vitro and an in vivo human:murine chimeric model. While APOBEC editors can reportedly exhibit guide-independent deamination of both DNA and RNA, we found no problematic 'off-target' activity or promiscuous base conversion affecting CAR antigen-specific binding regions, which may otherwise redirect T cell specificity. Combinational infusion of fratricide-resistant anti-T CAR T cells may enable enhanced molecular remission ahead of allo-HSCT for T cell malignancies.
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Developing an efficient and visible prime editing system to restore tobacco 8-hydroxy-copalyl diphosphate gene for labdane diterpene Z-abienol biosynthesis.
Prime editing (PE) is a versatile CRISPR-Cas based precise genome-editing platform widely used to introduce a range of possible base conversions in various organisms. However, no PE systems have been shown to induce heritable mutations in tobacco, nor in any other dicot. In this study, we generated an efficient PE system in tobacco that not only introduced heritable mutations, but also enabled anthocyanin-based reporter selection of transgene-free T1 plants. This system was used to confer Z-abienol biosynthesis in the allotetraploid tobacco cultivar HHDJY by restoring a G>T conversion in the NtCPS2 gene. High levels of Z-abienol were detected in the leaves of homozygous T1 plants at two weeks after topping. This study describes an advance in PE systems and expands genome-editing toolbox in tobacco, even in dicots, for use in basic research and molecular breeding. And restoring biosynthesis of Z-abienol in tobacco might provide an efficient way to obtain Z-abienol in plants.
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Genome Editing with CRISPR-Cas9: Can It Get Any Better?
The CRISPR-Cas revolution is taking place in virtually all fields of life sciences. Harnessing DNA cleavage with the CRISPR-Cas9 system of Streptococcus pyogenes has proven to be extraordinarily simple and efficient, relying only on the design of a synthetic single guide RNA (sgRNA) and its co-expression with Cas9. Here, we review the progress in the design of sgRNA from the original dual RNA guide for S. pyogenes and Staphylococcus aureus Cas9 (SpCas9 and SaCas9). New assays for genome-wide identification of off-targets have provided important insights into the issue of cleavage specificity in vivo. At the same time, the on-target activity of thousands of guides has been determined. These data have led to numerous online tools that facilitate the selection of guide RNAs in target sequences. It appears that for most basic research applications, cleavage activity can be maximized and off-targets minimized by carefully choosing guide RNAs based on computational predictions. Moreover, recent studies of Cas proteins have further improved the flexibility and precision of the CRISPR-Cas toolkit for genome editing. Inspired by the crystal structure of the complex of sgRNA-SpCas9 bound to target DNA, several variants of SpCas9 have recently been engineered, either with novel protospacer adjacent motifs (PAMs) or with drastically reduced off-targets. Novel Cas9 and Cas9-like proteins called Cpf1 have also been characterized from other bacteria and will benefit from the insights obtained from SpCas9. Genome editing with CRISPR-Cas9 may also progress with better understanding and control of cellular DNA repair pathways activated after Cas9-induced DNA cleavage.
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Improving the efficiency of precise genome editing with site-specific Cas9-oligonucleotide conjugates.
Site-specific chemical conjugation of proteins can enhance their therapeutic and diagnostic utility but has seldom been applied to CRISPR-Cas9, which is a rapidly growing field with great therapeutic potential. The low efficiency of homology-directed repair remains a major hurdle in CRISPR-Cas9-mediated precise genome editing, which is limited by low concentration of donor DNA template at the cleavage site. In this study, we have developed methodology to site-specifically conjugate oligonucleotides to recombinant Cas9 protein containing a genetically encoded noncanonical amino acid with orthogonal chemical reactivity. The Cas9-oligonucleotide conjugates recruited an unmodified donor DNA template to the target site through base pairing, markedly increasing homology-directed repair efficiency in both human cell culture and mouse zygotes. These chemically modified Cas9 mutants provide an additional tool, one that is complementary to chemically modified nucleic acids, for improving the utility of CRISPR-Cas9-based genome-editing systems.
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Hi-TOM: a platform for high-throughput tracking of mutations induced by CRISPR/Cas systems.
The CRISPR/Cas system has been extensively applied to make precise genetic modifications in various organisms. Despite its importance and widespread use, large-scale mutation screening remains time-consuming, labour-intensive and costly. Here, we developed Hi-TOM (available at https://doi.org/www.hi-tom.net/hi-tom/ ), an online tool to track the mutations with precise percentage for multiple samples and multiple target sites. We also described a corresponding next-generation sequencing (NGS) library construction strategy by fixing the bridge sequences and barcoding primers. Analysis of the samples from rice, hexaploid wheat and human cells reveals that the Hi-TOM tool has high reliability and sensitivity in tracking various mutations, especially complex chimeric mutations frequently induced by genome editing. Hi-TOM does not require special design of barcode primers, cumbersome parameter configuration or additional data analysis. Thus, the streamlined NGS library construction and comprehensive result output make Hi-TOM particularly suitable for high-throughput identification of all types of mutations induced by CRISPR/Cas systems.
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A Plug-in system for reprogramming the editing patterns of base editors.
DNA base editors are transformative genome editing tools that enable nucleotide conversions without inducing double-stranded DNA breaks, making them promising for correcting genetic mutations. Current base editors, however, are limited by fixed editing windows and constrained location of deaminases. To address these constraints, we develop a modular system termed Plug-in Base Editor (Plug-in BE), which dynamically programs deaminase positioning via integrating various epitopes and antibody-fused deaminases. This system expands the editing capabilities of base editors by optimizing deaminase's spatial interaction with DNA, leading to improvements in efficiencies, window restrictions, and safety profiles. We validate Plug-in BE's versatility and high fidelity in cancer gene therapy and zebrafish embryo editing, demonstrating its potential as a powerful and adaptable tool for basic research and therapeutic applications. This innovation can generate a series of base editors without extensive protein evolution, positioning Plug-in BE as a significant advancement in the field of genome editing.
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Structure-Guided Engineering of a Cas12i Nuclease Unlocks Near-PAMless Genome Editing.
The therapeutic and research applications of CRISPR-Cas nucleases are constrained by their reliance on specific Protospacer Adjacent Motifs (PAMs), which limit the accessible sites in the genome. To overcome this critical barrier, we performed structure-guided engineering of SF01, a compact Cas12i nuclease. Using AlphaFold-predicted structural models, we identified and systematically mutagenized 38 residues at the PAM-interacting interface. This iterative engineering process yielded three superior variants-KR, IKRR, and STKRR-that exhibit dramatically relaxed PAM specificity, enabling efficient editing at a broad spectrum of 5'-NNTN-3' sites. Importantly, while the most broad-spectrum variant (STKRR) shows a trade-off at canonical sites, the IKRR variant retains high activity at canonical 5'-NTTN-3' PAMs while simultaneously enabling efficient editing at 5'-NNTN-3' sites. This near-PAMless activity expands the targetable portion of the genome to over 25%, a four-fold increase over the parental nuclease. Furthermore, adenine base editors (ABEs) constructed with these variants achieve high-efficiency editing (∼80%) at endogenous loci with expanded targeting scope. Comprehensive off-target analysis using GUIDE-tag and Digenome-seq revealed that the enhanced on-target activity of the SF01 variants is not accompanied by a loss of specificity. These engineered nucleases represent a powerful and versatile expansion of the genome editing toolkit, enabling applications previously inaccessible due to PAM constraints.
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Programmable deletion, replacement, integration and inversion of large DNA sequences with twin prime editing.
The targeted deletion, replacement, integration or inversion of genomic sequences could be used to study or treat human genetic diseases, but existing methods typically require double-strand DNA breaks (DSBs) that lead to undesired consequences, including uncontrolled indel mixtures and chromosomal abnormalities. Here we describe twin prime editing (twinPE), a DSB-independent method that uses a prime editor protein and two prime editing guide RNAs (pegRNAs) for the programmable replacement or excision of DNA sequences at endogenous human genomic sites. The two pegRNAs template the synthesis of complementary DNA flaps on opposing strands of genomic DNA, which replace the endogenous DNA sequence between the prime-editor-induced nick sites. When combined with a site-specific serine recombinase, twinPE enabled targeted integration of gene-sized DNA plasmids (>5,000 bp) and targeted sequence inversions of 40 kb in human cells. TwinPE expands the capabilities of precision gene editing and might synergize with other tools for the correction or complementation of large or complex human pathogenic alleles.
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Implementing the CRISPR/Cas9 Technology in Eucalyptus Hairy Roots Using Wood-Related Genes.
Eucalypts are the most planted hardwoods worldwide. The availability of the Eucalyptus grandis genome highlighted many genes awaiting functional characterization, lagging behind because of the lack of efficient genetic transformation protocols. In order to efficiently generate knock-out mutants to study the function of eucalypts genes, we implemented the powerful CRISPR/Cas9 gene editing technology with the hairy roots transformation system. As proofs-of-concept, we targeted two wood-related genes: Cinnamoyl-CoA Reductase1 (CCR1), a key lignin biosynthetic gene and IAA9A an auxin dependent transcription factor of Aux/IAA family. Almost all transgenic hairy roots were edited but the allele-editing rates and spectra varied greatly depending on the gene targeted. Most edition events generated truncated proteins, the prevalent edition types were small deletions but large deletions were also quite frequent. By using a combination of FT-IR spectroscopy and multivariate analysis (partial least square analysis (PLS-DA)), we showed that the CCR1-edited lines, which were clearly separated from the controls. The most discriminant wave-numbers were attributed to lignin. Histochemical analyses further confirmed the decreased lignification and the presence of collapsed vessels in CCR1-edited lines, which are characteristics of CCR1 deficiency. Although the efficiency of editing could be improved, the method described here is already a powerful tool to functionally characterize eucalypts genes for both basic research and industry purposes.
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