CRISPR Nucleases Research Articles

CRISPR-Cas9/Cas12a systems for efficient genome editing and large genomic fragment deletions in Aspergillus niger.

CRISPR technology has revolutionized fungal genetic engineering by accelerating the pace and expanding the feasible scope of experiments in this field. Among various CRISPR-Cas systems, Cas9 and Cas12a are widely used in genetic and metabolic engineering. In filamentous fungi, both Cas9 and Cas12a have been utilized as CRISPR nucleases. In this work we first compared efficacies and types of genetic edits for CRISPR-Cas9 and -Cas12a systems at the polyketide synthase (albA) gene locus in Aspergillus niger. By employing a tRNA-based gRNA polycistronic cassette, both Cas9 and Cas12a have demonstrated equally remarkable editing efficacy. Cas12a showed potential superiority over Cas9 protein when one gRNA was used for targeting, achieving an editing efficiency of 86.5% compared to 31.7% for Cas9. Moreover, when employing two gRNAs for targeting, both systems achieved up to 100% editing efficiency for single gene editing. In addition, the CRISPR-Cas9 system has been reported to induce large genomic deletions in various species. However, its use for engineering large chromosomal segments deletions in filamentous fungi still requires optimization. Here, we engineered Cas9 and -Cas12a-induced large genomic fragment deletions by targeting various genomic regions of A. niger ranging from 3.5kb to 40kb. Our findings demonstrate that targeted engineering of large chromosomal segments can be achieved, with deletions of up to 69.1% efficiency. Furthermore, by targeting a secondary metabolite gene cluster, we show that fragments over 100kb can be efficiently and specifically deleted using the CRISPR-Cas9 or -Cas12a system. Overall, in this paper, we present an efficient multi-gRNA genome editing system utilizing Cas9 or Cas12a that enables highly efficient targeted editing of genes and large chromosomal regions in A. niger.

High-fidelity PAMless base editing of hematopoietic stem cells to treat chronic granulomatous disease.

X-linked chronic granulomatous disease (X-CGD) is an inborn error of immunity (IEI) resulting from genetic mutations in the cytochrome b-245 beta chain (CYBB) gene. The applicability of base editors (BEs) to correct mutations that cause X-CGD is constrained by the requirement of Cas enzymes to recognize specific protospacer adjacent motifs (PAMs). Our recently engineered PAMless Cas enzyme, SpRY, can overcome the PAM limitation. However, the efficiency, specificity, and applicability of SpRY-based BEs to correct mutations in human hematopoietic stem and progenitor cells (HSPCs) have not been thoroughly examined. Here, we demonstrated that the adenine BE ABE8e-SpRY can access a range of target sites in HSPCs to correct mutations causative of X-CGD. For the prototypical X-CGD mutation CYBB c.676C>T, ABE8e-SpRY achieved up to 70% correction, reaching efficiencies greater than three-and-one-half times higher than previous CRISPR nuclease and donor template approaches. We profiled potential off-target DNA edits, transcriptome-wide RNA edits, and chromosomal perturbations in base-edited HSPCs, which together revealed minimal off-target or bystander edits. Edited alleles persisted after transplantation of the base-edited HSPCs into immunodeficient mice. Together, these investigational new drug-enabling studies demonstrated efficient and precise correction of an X-CGD mutation with PAMless BEs, supporting a first-in-human clinical trial (NCT06325709) and providing a potential blueprint for treatment of other IEI mutations.

AAV-mediated gene therapies by miniature gene editing tools.

The advent of CRISPR-Cas has revolutionized precise gene editing. While pioneering CRISPR nucleases like Cas9 and Cas12 generate targeted DNA double-strand breaks (DSB) for knockout or homology-directed repair, next generation CRISPR technologies enable gene editing without DNA DSB. Base editors directly convert bases, prime editors make diverse alterations, and dead Cas-regulator fusions allow nuanced control of gene expression, avoiding potentially risks like translocations. Meanwhile, the discovery of diminutive Cas12 orthologs and Obligate Mobile Element-Guided Activity (OMEGA) nucleases has overcome cargo limitations of adeno-associated viral vectors, expanding prospects for in vivo therapeutic delivery. Here, we review the ever-evolving landscape of cutting-edge gene editing tools, focusing on miniature Cas12 orthologs and OMEGA effectors amenable to single AAV packaging. We also summarize CRISPR therapies delivered using AAV vectors, discuss challenges such as efficiency and specificity, and look to the future of this transformative field of in vivo gene editing enabled by AAV vectors delivery.

CRISPRoff epigenetic editing for programmable gene silencing in human cells without DNA breaks.

The advent of CRISPR-based technologies has enabled the rapid advancement of programmable gene manipulation in cells, tissues, and whole organisms. An emerging platform for targeted gene perturbation is epigenetic editing, the direct editing of chemical modifications on DNA and histones that ultimately results in repression or activation of the targeted gene. In contrast to CRISPR nucleases, epigenetic editors modulate gene expression without inducing DNA breaks or altering the genomic sequence of host cells. Recently, we developed the CRISPRoff epigenetic editing technology that simultaneously establishes DNA methylation and repressive histone modifications at targeted gene promoters. Transient expression of CRISPRoff and the accompanying single guide RNAs in mammalian cells results in transcriptional repression of targeted genes that is memorized heritably by cells through cell division and differentiation. Here, we describe our protocol for the delivery of CRISPRoff through plasmid DNA transfection, as well as the delivery of CRISPRoff mRNA, into transformed human cell lines and primary immune cells. We also provide guidance on evaluating target gene silencing and highlight key considerations when utilizing CRISPRoff for gene perturbations. Our protocols are broadly applicable to other CRISPR-based epigenetic editing technologies, as programmable genome manipulation tools continue to evolve rapidly.

PSLBII-12 Cloning and analysis of target-specific CRISPR RNA and screening primers to create β-lactoglobulin knock-out goat fibroblast cell lines for nuclear transfer

Abstract Goat milk and meat are primary or additional source of income for small farmers in large parts of the world. Goat farming is increasing in southeastern United States, due to influx of ethnic populations. Goat milk is rich in health promoting nutrients compared with other livestock milk; however, its consumption results in allergenicity in humans due to presence of β-lactoglobulin protein in milk. Although some efforts have been made to eliminate allergenicity by modifying β-lactoglobulin gene, especially in Chinese goats, more research is needed to make it a reality. The objective of this study was to design, clone and characterize CRISPR RNA (crRNA) in a plasmid vector containing Cas 9 expression cassette as well as design and analyze PCR primers for efficient screening of edited genomes as a prelude to create and analyze β-lactoglobulin knock-out goats. Three crRNAs, two from exon 1 and one from exon 2 of goat β-lactoglobulin gene with 5 base hangovers were designed and cloned individually in GeneArt CRISPR Nuclease plasmid Vector. Sequence analysis of selected plasmid clones using U 6 promoter primer, upstream of the crRNA insert, confirmed that at least 3 clones of each crRNA had the right sequence. Additionally, 4 sets (forward and reverse) of primers to amplify 4 loci in β-lactoglobulin gene flanking the crRNA sequences in goat genome were designed. PCR amplification using genomic DNA isolated from goat blood as well as goat skin fibroblasts (GSF) confirmed expected single bands of 655, 812, 502 and 437 bps with respective primer sets. Transfection of 3 crRNA clones in GSF cells and their analysis for site-specific genome modification is in progress. In conclusion, we have designed, cloned, and tested crRNA in a plasmid vector along with 4 sets of PCR primers to screen for edited genomes which will be useful to create β-lactoglobulin edited goats to eliminate milk allergenicity.

Rapid protein evolution by few-shot learning with a protein language model.

Directed evolution of proteins is critical for applications in basic biological research, therapeutics, diagnostics, and sustainability. However, directed evolution methods are labor intensive, cannot efficiently optimize over multiple protein properties, and are often trapped by local maxima. In silico- directed evolution methods incorporating protein language models (PLMs) have the potential to accelerate this engineering process, but current approaches fail to generalize across diverse protein families. We introduce EVOLVEpro, a few-shot active learning framework to rapidly improve protein activity using a combination of PLMs and protein activity predictors, achieving improved activity with as few as four rounds of evolution. EVOLVEpro substantially enhances the efficiency and effectiveness of in silico protein evolution, surpassing current state-of-the-art methods and yielding proteins with up to 100-fold improvement of desired properties. We showcase EVOLVEpro for five proteins across three applications: T7 RNA polymerase for RNA production, a miniature CRISPR nuclease, a prime editor, and an integrase for genome editing, and a monoclonal antibody for epitope binding. These results demonstrate the advantages of few-shot active learning with small amounts of experimental data over zero-shot predictions. EVOLVEpro paves the way for broader applications of AI-guided protein engineering in biology and medicine.

CRISPR-based gene editing in plants: Focus on reagents and their delivery tools

Introduction: CRISPR-Cas9 technology has revolutionized plant genome editing, providing precise and efficient methods for genetic modification. This study focuses on the advancements and delivery of CRISPR-Cas9 in plant gene editing. Methods: A comprehensive search in scientific databases, including PubMed, ScienceDirect, and Google Scholar, was conducted to gather information on CRISPR-Cas9 gene editing and its delivery in precise gene modification in plants. Results: The evolving landscape of CRISPR nucleases has led to the development of innovative technologies, enhancing plant research. However, successful editing is contingent on efficient delivery of genome engineering reagents. CRISPR-based gene editing in plants utilizes diverse delivery methods: Agrobacterium-mediated transformation for bacterial transfer, biolistic transformation for physical gene insertion, electroporation for direct gene entry, expression of developmental regulators for gene expression modulation, and tobacco rattle virus as a viral vector, each offering distinct advantages for precise and efficient genetic modification in plants. Conclusion: CRISPR-Cas9 gene editing stands as a pivotal advancement in plant genetics, offering precise gene manipulation with applications in agriculture and biotechnology. The continuous refinement of reagent delivery tools reinforces CRISPR-Cas9's transformative role in plant genome editing, with significant implications for broader scientific applications.

Systematic interrogation of CRISPR antimicrobials in Klebsiella pneumoniae reveals nuclease-, guide-and strain-dependent features influencing antimicrobial activity.

CRISPR-Cas systems can be utilized as programmable-spectrum antimicrobials to combat bacterial infections. However, how CRISPR nucleases perform as antimicrobials across target sites and strains remains poorly explored. Here, we address this knowledge gap by systematically interrogating the use of CRISPR antimicrobials using multidrug-resistant and hypervirulent strains of Klebsiella pneumoniae as models. Comparing different Cas nucleases, DNA-targeting nucleases outperformed RNA-targeting nucleases based on the tested targets. Focusing on AsCas12a that exhibited robust targeting across different strains, we found that the elucidated modes of escape varied widely, restraining opportunities to enhance killing. We also encountered individual guide RNAs yielding different extents of clearance across strains, which were linked to an interplay between improper gRNA folding and strain-specific DNA repair and survival. To explore features that could improve targeting across strains, we performed a genome-wide screen in different K. pneumoniae strains that yielded guide design rules and trained an algorithm for predicting guide efficiency. Finally, we showed that Cas12a antimicrobials can be exploited to eliminate K. pneumoniae when encoded in phagemids delivered by T7-like phages. Altogether, our results highlight the importance of evaluating antimicrobial activity of CRISPR antimicrobials across relevant strains and define critical parameters for efficient CRISPR-based targeting.

CRISPR-Cas-mediated Multianalyte Synthetic Urine Biomarker Test for Portable Diagnostics.

Creating synthetic biomarkers for the development of precision diagnostics has enabled detection of disease through pathways beyond those used for traditional biofluid measurements. Synthetic biomarkers generally make use of reporters that provide readable signals in the biofluid to reflect the biochemical alterations in the local disease microenvironment during disease incidence and progression. The pharmacokinetic concentration of the reporters and biochemical amplification of the disease signal are paramount to achieving high sensitivity and specificity in a diagnostic test. Here, a cancer diagnostic platform is built using one format of synthetic biomarkers: activity-based nanosensors carrying chemically stabilized DNA reporters that can be liberated by aberrant proteolytic signatures in the tumor microenvironment. Synthetic DNA as a disease reporter affords multiplexing capability through its use as a barcode, allowing for the readout of multiple proteolytic signatures at once. DNA reporters released into the urine are detected using CRISPR nucleases via hybridization with CRISPR RNAs, which in turn produce a fluorescent or colorimetric signal upon enzyme activation. In this protocol, DNA-barcoded, activity-based nanosensors are constructed and their application is exemplified in a preclinical mouse model of metastatic colorectal cancer. This system is highly modifiable according to disease biology and generates multiple disease signals simultaneously, affording a comprehensive understanding of the disease characteristics through a minimally invasive process requiring only nanosensor administration, urine collection, and a paper test which enables point-of-care diagnostics.

A temperature-tolerant CRISPR base editor mediates highly efficient and precise gene editing in Drosophila.

CRISPR nucleases generate a broad spectrum of mutations that includes undesired editing outcomes. Here, we develop optimized C-to-T base editing systems for the generation of precise loss- or gain-of-function alleles in Drosophila and identify temperature as a crucial parameter for efficiency. We find that a variant of the widely used APOBEC1 deaminase has attenuated activity at 18° to 29°C and shows considerable dose-dependent toxicity. In contrast, the temperature-tolerant evoCDA1 domain mediates editing of typically more than 90% of alleles and is substantially better tolerated. Furthermore, formation of undesired mutations is exceptionally rare in Drosophila compared to other species. The predictable editing outcome, high efficiency, and product purity enables near homogeneous induction of STOP codons or alleles encoding protein variants in vivo. Last, we demonstrate how optimized expression enables conditional base editing in marked cell populations. This work substantially facilitates creation of precise alleles in Drosophila and provides key design parameters for developing efficient base editing systems in other ectothermic species.

The Possibilities of Gene Drives for Managing Populations and Controlling Diseases

The technical limitations and the use of gene drives to address ecological problems by modifying all populations of wild species remain primarily speculative. Here, we examine the possibility that RNA-guided gene drives based on the CRISPR nuclease Cas9 could be used as an all-encompassing approach for introducing changed features into natural populations over a long period. We outline potential capabilities and possible disadvantages and offer new preventative measures to stem from the propagation of genes and undo genetic modifications. Editing the sexual animal population would significantly benefit both people and the environment. For instance, RNA-guided gene drives may stop the spread of illness, assist farming by correcting bug and weed chemicals and resistance to herbicides, and manage harmful invasive species. However, each prospective use needs to be carefully evaluated due to the likelihood of unfavorable ecological repercussions and the near inevitability of dissemination beyond political boundaries. To investigate the responsible application of this now hypothetical technology, we want serious, inclusive, educated public conversations

CRISPR-based diagnostics of different biomolecules from nucleic acids, proteins, and small molecules to exosomes.

CRISPR-based detection technologies have been widely explored for molecular diagnostics. However, the challenge lies in converting the signal of different biomolecules, such as nucleic acids, proteins, small molecules, exosomes, and ions, into a CRISPR-based nucleic acid detection signal. Understanding the detection of different biomolecules using CRISPR technology can aid in the development of practical and promising detection approaches. Unfortunately, existing reviews rarely provide an overview of CRISPR-based molecular diagnostics from the perspective of different biomolecules. Herein, we first introduce the principles and characteristics of various CRISPR nucleases for molecular diagnostics. Then, we focus on summarizing and evaluating the latest advancements in CRISPR-based detection of different biomolecules. Through a comparison of different methods of amplification and signal readout, we discuss how general detection methods can be integrated with CRISPR. Finally, we conclude by identifying opportunities for the improvement of CRISPR in quantitative, amplification-free, multiplex, all-in-one, and point-of-care testing (POCT) purposes.

A massively parallel approach for assessing CRISPR off-targets in vitro

In a recent issue of Med, Tian etal.1 present AID-seq, an approach that enables massively parallel identification of off-targets for different CRISPR nucleases invitro. By using a pooled strategy to simultaneously identify the on-/off-targets of multiple gRNAs, the authors could screen the most efficient and safe gRNA candidates.

Dual-locus, dual-HDR editing permits efficient generation of antigen-specific regulatory T cells with robust suppressive activity

Adoptive regulatory T (Treg) cell therapy is predicted to modulate immune tolerance in autoimmune diseases, including type 1 diabetes (T1D). However, the requirement for antigen (ag) specificity to optimally orchestrate tissue-specific, Treg cell-mediated tolerance limits effective clinical application. To address this challenge, we present a single-step, combinatorial gene editing strategy utilizing dual-locus, dual-homology-directed repair (HDR) to generate and specifically expand ag-specific engineered Treg (EngTreg) cells derived from donor CD4+ Tcells. Concurrent delivery of CRISPR nucleases and recombinant (r)AAV homology donor templates targeting FOXP3 and TRAC was used to achieve three parallel goals: enforced, stable expression of FOXP3; replacement of the endogenous Tcell receptor (TCR) with an islet-specific TCR; and selective enrichment of dual-edited cells. Each HDR donor template contained an alternative component of a heterodimeric chemically inducible signaling complex (CISC), designed to activate interleukin-2 (IL-2) signaling in response to rapamycin, promoting expansion of only dual-edited EngTreg cells. Using this approach, we generated purified, islet-specific EngTreg cells that mediated robust direct and bystander suppression of effector T (Teff) cells recognizing the same or a different islet antigen peptide, respectively. This platform is broadly adaptable for use with alternative TCRs or other targeting moieties for application in tissue-specific autoimmune or inflammatory diseases.

Cellular Surveillance: DNA-Based Recording to Monitor and Memorize Biological Events

Continuous monitoring of intra- and extracellular environments, ranging from open fields and rivers to within a single cell, is often limited by costly equipment, labor intensity, or invasive techniques. It may, however, be possible to avoid these issues by developing a monitoring device on a much smaller scale. By applying novel synthetic biology techniques, it is now possible to engineer organisms with refined biorecording capabilities, using DNA as a recording medium on which a cellular “memory” of past events can be written. Through a variety of mechanisms, using recombinases, CRISPR nucleases, base editing, polymerases, and prime editing, intra- and extracellular events can be recorded along with temporal, spatial, and magnitude information. Here, the mechanisms developed for DNA-based biorecording devices so far are assessed and discussed, within the context of the potential future applications of these devices.

A highly efficient transgene knock-in technology in clinically relevant cell types.

Inefficient knock-in of transgene cargos limits the potential of cell-based medicines. In this study, we used a CRISPR nuclease that targets a site within an exon of an essential gene and designed a cargo template so that correct knock-in would retain essential gene function while also integrating the transgene(s) of interest. Cells with non-productive insertions and deletions would undergo negative selection. This technology, called SLEEK (SeLection by Essential-gene Exon Knock-in), achieved knock-in efficiencies of more than 90% in clinically relevant cell types without impacting long-term viability or expansion. SLEEK knock-in rates in T cells are more efficient than state-of-the-art TRAC knock-in with AAV6 and surpass more than 90% efficiency even with non-viral DNA cargos. As a clinical application, natural killer cells generated from induced pluripotent stem cells containing SLEEK knock-in of CD16 and mbIL-15 show substantially improved tumor killing and persistence in vivo.

Boosting genome editing efficiency in human cells and plants with novel LbCas12a variants

BackgroundCas12a (formerly known as Cpf1), the class II type V CRISPR nuclease, has been widely used for genome editing in mammalian cells and plants due to its distinct characteristics from Cas9. Despite being one of the most robust Cas12a nucleases, LbCas12a in general is less efficient than SpCas9 for genome editing in human cells, animals, and plants.ResultsTo improve the editing efficiency of LbCas12a, we conduct saturation mutagenesis in E. coli and identify 1977 positive point mutations of LbCas12a. We selectively assess the editing efficiency of 56 LbCas12a variants in human cells, identifying an optimal LbCas12a variant (RVQ: G146R/R182V/E795Q) with the most robust editing activity. We further test LbCas12a-RV, LbCas12a-RRV, and LbCas12a-RVQ in plants and find LbCas12a-RV has robust editing activity in rice and tomato protoplasts. Interestingly, LbCas12a-RRV, resulting from the stacking of RV and D156R, displays improved editing efficiency in stably transformed rice and poplar plants, leading to up to 100% editing efficiency in T0 plants of both plant species. Moreover, this high-efficiency editing occurs even at the non-canonical TTV PAM sites.ConclusionsOur results demonstrate that LbCas12a-RVQ is a powerful tool for genome editing in human cells while LbCas12a-RRV confers robust genome editing in plants. Our study reveals the tremendous potential of these LbCas12a variants for advancing precision genome editing applications across a wide range of organisms.

CRISPR-Cas-amplified urinary biomarkers for multiplexed and portable cancer diagnostics

Synthetic biomarkers, bioengineered sensors that generate molecular reporters in diseased microenvironments, represent an emerging paradigm in precision diagnostics. Despite the utility of DNA barcodes as a multiplexing tool, their susceptibility to nucleases in vivo has limited their utility. Here we exploit chemically stabilized nucleic acids to multiplex synthetic biomarkers and produce diagnostic signals in biofluids that can be ‘read out’ via CRISPR nucleases. The strategy relies on microenvironmental endopeptidase to trigger the release of nucleic acid barcodes and polymerase-amplification-free, CRISPR-Cas-mediated barcode detection in unprocessed urine. Our data suggest that DNA-encoded nanosensors can non-invasively detect and differentiate disease states in transplanted and autochthonous murine cancer models. We also demonstrate that CRISPR-Cas amplification can be harnessed to convert the readout to a point-of-care paper diagnostic tool. Finally, we employ a microfluidic platform for densely multiplexed, CRISPR-mediated DNA barcode readout that can potentially evaluate complex human diseases rapidly and guide therapeutic decisions.

Combining different CRISPR nucleases for simultaneous knock-in and base editing prevents translocations in multiplex-edited CAR T cells

BackgroundMultiple genetic modifications may be required to develop potent off-the-shelf chimeric antigen receptor (CAR) T cell therapies. Conventional CRISPR-Cas nucleases install sequence-specific DNA double-strand breaks (DSBs), enabling gene knock-out or targeted transgene knock-in. However, simultaneous DSBs provoke a high rate of genomic rearrangements which may impede the safety of the edited cells.ResultsHere, we combine a non-viral CRISPR-Cas9 nuclease-assisted knock-in and Cas9-derived base editing technology for DSB free knock-outs within a single intervention. We demonstrate efficient insertion of a CAR into the T cell receptor alpha constant (TRAC) gene, along with two knock-outs that silence major histocompatibility complexes (MHC) class I and II expression. This approach reduces translocations to 1.4% of edited cells. Small insertions and deletions at the base editing target sites indicate guide RNA exchange between the editors. This is overcome by using CRISPR enzymes of distinct evolutionary origins. Combining Cas12a Ultra for CAR knock-in and a Cas9-derived base editor enables the efficient generation of triple-edited CAR T cells with a translocation frequency comparable to unedited T cells. Resulting TCR- and MHC-negative CAR T cells resist allogeneic T cell targeting in vitro.ConclusionsWe outline a solution for non-viral CAR gene transfer and efficient gene silencing using different CRISPR enzymes for knock-in and base editing to prevent translocations. This single-step procedure may enable safer multiplex-edited cell products and demonstrates a path towards off-the-shelf CAR therapeutics.

Anzalone Prime: An Interview with Prime Editing Developer Andrew Anzalone

Anzalone Prime: An Interview with Prime Editing Developer Andrew Anzalone