RNA editing catalytic complexes edit multiple mRNA sites non-processively in Trypanosoma brucei.

RNA editing generates mature mitochondrial mRNAs in T. brucei by extensive uridine insertion and deletion at numerous editing sites (ESs) as specified by guide RNAs (gRNAs). The editing is performed by three RNA Editing Catalytic Complexes (RECCs) which each have a different endonuclease in addition to 12 proteins in common resulting in RECC1 that is specific for deletion ESs and RECC2 and RECC3 that are specific for insertion ESs. Thus, different RECCs are required for editing of mRNA sequence regions where single gRNAs specify a combination of insertion and deletion ESs. We investigated how the three different RECCs might edit combinations of insertion and deletion ESs that are specified by single gRNAs by testing whether their endonuclease compositions are stable or dynamic during editing. We analyzed in vivo BirA* proximity labeling and found that the endonucleases remain associated with their set of common RECC proteins during editing when expressed at normal physiological levels. We also found that overexpression of endonuclease components resulted in minor effects on RECCs but did not affect growth. Thus, the protein stoichiometries that exist within each RECC can be altered by perturbations of RECC expression levels. These results indicate that editing of consecutive insertion and deletion ESs occurs by successive engagement and disengagement of RECCs, i.e., is non-processive, which is likely the case for consecutive pairs of insertion or deletion ESs. This clarifies the nature of the complex patterns of partially edited mRNAs that occur in vivo.

Structural basis for guide RNA selection by the RESC1-RESC2 complex.

Kinetoplastid parasites, such as trypanosomes or leishmania, rely on RNA-templated RNA editing to mature mitochondrial cryptic pre-mRNAs into functional protein-coding transcripts. Processive pan-editing of multiple editing blocks within a single transcript is dependent on the 20-subunit RNA editing substrate binding complex (RESC) that serves as a platform to orchestrate the interactions between pre-mRNA, guide RNAs (gRNAs), the catalytic RNA editing complex (RECC), and a set of RNA helicases. Due to the lack of molecular structures and biochemical studies with purified components, neither the spacio-temporal interplay of these factors nor the selection mechanism for the different RNA components is understood. Here we report the cryo-EM structure of Trypanosoma brucei RESC1-RESC2, a central hub module of the RESC complex. The structure reveals that RESC1 and RESC2 form an obligatory domain-swapped dimer. Although the tertiary structures of both subunits closely resemble each other, only RESC2 selectively binds 5'-triphosphate-nucleosides, a defining characteristic of gRNAs. We therefore propose RESC2 as the protective 5'-end binding site for gRNAs within the RESC complex. Overall, our structure provides a starting point for the study of the assembly and function of larger RNA-bound kinetoplast RNA editing modules and might aid in the design of anti-parasite drugs.

Structure of the T. brucei kinetoplastid RNA editing substrate-binding complex core component, RESC5.

Kinetoplastid protists such as Trypanosoma brucei undergo an unusual process of mitochondrial uridine (U) insertion and deletion editing termed kinetoplastid RNA editing (kRNA editing). This extensive form of editing, which is mediated by guide RNAs (gRNAs), can involve the insertion of hundreds of Us and deletion of tens of Us to form a functional mitochondrial mRNA transcript. kRNA editing is catalyzed by the 20 S editosome/RECC. However, gRNA directed, processive editing requires the RNA editing substrate binding complex (RESC), which is comprised of 6 core proteins, RESC1-RESC6. To date there are no structures of RESC proteins or complexes and because RESC proteins show no homology to proteins of known structure, their molecular architecture remains unknown. RESC5 is a key core component in forming the foundation of the RESC complex. To gain insight into the RESC5 protein we performed biochemical and structural studies. We show that RESC5 is monomeric and we report the T. brucei RESC5 crystal structure to 1.95 Å. RESC5 harbors a dimethylarginine dimethylaminohydrolase-like (DDAH) fold. DDAH enzymes hydrolyze methylated arginine residues produced during protein degradation. However, RESC5 is missing two key catalytic DDAH residues and does bind DDAH substrate or product. Implications of the fold for RESC5 function are discussed. This structure provides the first structural view of an RESC protein.

KRGG1 function in RNA editing in Trypanosoma brucei.

Mitochondrial gene expression in trypanosomes requires numerous multiprotein complexes that are unique to kinetoplastids. Among these, the most well characterized are RNA editing catalytic complexes (RECCs) that catalyze the guide RNA (gRNA)-specified insertion and deletion of uridines during mitochondrial mRNA maturation. This post-transcriptional resequencing of mitochondrial mRNAs can be extensive, involving dozens of different gRNAs and hundreds of editing sites with most of the mature mRNA sequences resulting from the editing process. Proper coordination of the editing with the cognate gRNAs is attributed to RNA editing substrate-binding complexes (RESCs), which are also required for RNA editing. Although the precise mechanism of RESC function is less well understood, their affinity for binding both editing substrates and products suggests that these complexes may provide a scaffold for RECC catalytic processing. KRGG1 has been shown to bind RNAs, and although affinity purification co-isolates RESC complexes, its role in RNA editing remains uncertain. We show here that KRGG1 is essential in BF parasites and required for normal editing. KRGG1 repression results in reduced amounts of edited A6 mRNA and increased amounts of edited ND8 mRNA. Sequence and structure analysis of KRGG1 identified a region of homology with RESC6, and both proteins have predicted tandem helical repeats that resemble ARM/HEAT motifs. The ARM/HEAT-like region is critical for function as exclusive expression of mutated KRGG1 results in growth inhibition and disruption of KRGG1 association with RESCs. These results indicate that KRGG1 is critical for RNA editing and its specific function is associated with RESC activity.

Improving Precise Genome Editing Using Donor DNA/gRNA Hybrid Duplex Generated by Complementary Bases.

In precise genome editing, site-specific DNA double-strand breaks (DSBs) induced by the CRISPR/Cas9 system are repaired via homology-directed repair (HDR) using exogenous donor DNA templates. However, the low efficiency of HDR-mediated genome editing is a barrier to widespread use. In this study, we created a donor DNA/guide RNA (gRNA) hybrid duplex (DGybrid) that was composed of sequence-extended gRNA and single-stranded oligodeoxynucleotide (ssODN) combined with complementary bases without chemical modifications to increase the concentration of donor DNA at the cleavage site. The efficiency of genome editing using DGybrid was evaluated in Saccharomyces cerevisiae. The results show a 1.8-fold (from 35% to 62%) improvement in HDR-mediated editing efficiency compared to genome editing in which gRNA and donor DNA were introduced separately. In addition, analysis of the nucleic acid introduction efficiency using flow cytometry indicated that both RNA and ssODNs are efficiently incorporated into cells together by using the DNA/RNA hybrid. Our technique would be preferred as a universal and concise tool for improving the efficiency of HDR-mediated genome editing.

Self-cleaving guide RNAs enable pharmacological selection of precise gene editing events in vivo.

Expression of guide RNAs in the CRISPR/Cas9 system typically requires the use of RNA polymerase III promoters, which are not cell-type specific. Flanking the gRNA with self-cleaving ribozyme motifs to create a self-cleaving gRNA overcomes this limitation. Here, we use self-cleaving gRNAs to create drug-selectable gene editing events in specific hepatocyte loci. A recombinant Adeno Associated Virus vector targeting the Albumin locus with a promoterless self-cleaving gRNA to create drug resistance is linked in cis with the therapeutic transgene. Gene expression of both are dependent on homologous recombination into the target locus. In vivo drug selection for the precisely edited hepatocytes allows >30-fold expansion of gene-edited cells and results in therapeutic levels of a human Factor 9 transgene. Importantly, self-cleaving gRNA expression is also achieved after targeting weak hepatocyte genes. We conclude that self-cleaving gRNAs are a powerful system to enable cell-type specific in vivo drug resistance for therapeutic gene editing applications.

New Effective Method of Lactococcus Genome Editing Using Guide RNA-Directed Transposition.

Lactococcus lactis is an important industrial microorganism and a widely used model object for research in the field of lactic acid bacteria (LAB) biology. The development of new L. lactis and related LAB strains with improved properties, including phage-resistant strains for dairy fermentation, LAB-based vaccines or strains with altered genotypes for research purposes, are hindered by the lack of genome-editing tools that allow for the easy and straightforward incorporation of a significant amount of the novel genetic material, such as large genes or operons, into the chromosomes of these bacteria. We recently employed a suggested system based on the CRISPR-Cas-associated transposon for the editing of the L. lactis genome. After the in-depth redesign of the system, we were able to achieve the stable incorporation of the fragments that were sized up to 10 kbp into the L. lactis beta-galactosidase gene. The efficiency of editing under the optimized conditions were 2 × 10-4 and 4 × 10-5 for 1 kbp and 10 kbp, respectively, which are sufficient for fast and easy modifications if a positive selection marker can be used.

Antibody-mediated delivery of CRISPR-Cas9 ribonucleoproteins in human cells.

The CRISPR genome editing technology holds great clinical potential for the treatment of monogenetic disorders such as sickle cell disease. The therapeutic in vivo application of the technology relies on targeted delivery methods of the Cas9 and gRNA complex to specific cells or tissues. However, such methods are currently limited to direct organ delivery, preventing clinical application. Here, we show that monoclonal antibodies can be employed to deliver the Cas9/gRNA complex directly into human cells via cell-surface receptors. Using the SpyCatcher/SpyTag system, we conjugated the Fab fragment of the therapeutic antibodies Trastuzumab and Pertuzumab directly to the Cas9 enzyme and observed HER2-specific uptake of the ribonucleoprotein in a human HER2 expressing cell line. Following cellular uptake in the presence of an endosomolytic peptide, modest gene editing was also observed. This finding provides a blueprint for the targeted delivery of the CRISPR technology into specific cells using monoclonal antibodies.

A comprehensive Bioconductor ecosystem for the design of CRISPR guide RNAs across nucleases and technologies.

The success of CRISPR-mediated gene perturbation studies is highly dependent on the quality of gRNAs, and several tools have been developed to enable optimal gRNA design. However, these tools are not all adaptable to the latest CRISPR modalities or nucleases, nor do they offer comprehensive annotation methods for advanced CRISPR applications. Here, we present a new ecosystem of R packages, called crisprVerse, that enables efficient gRNA design and annotation for a multitude of CRISPR technologies. This includes CRISPR knockout (CRISPRko), CRISPR activation (CRISPRa), CRISPR interference (CRISPRi), CRISPR base editing (CRISPRbe) and CRISPR knockdown (CRISPRkd). The core package, crisprDesign, offers a user-friendly and unified interface to add off-target annotations, rich gene and SNP annotations, and on- and off-target activity scores. These functionalities are enabled for any RNA- or DNA-targeting nucleases, including Cas9, Cas12, and Cas13. The crisprVerse ecosystem is open-source and deployed through the Bioconductor project ( https://github.com/crisprVerse ).

Genome-wide base editor screen identifies regulators of protein abundance in yeast.

Proteins are key molecular players in a cell, and their abundance is extensively regulated not just at the level of gene expression but also post-transcriptionally. Here, we describe a genetic screen in yeast that enables systematic characterization of how protein abundance regulation is encoded in the genome. The screen combines a CRISPR/Cas9 base editor to introduce point mutations with fluorescent tagging of endogenous proteins to facilitate a flow-cytometric readout. We first benchmarked base editor performance in yeast with individual gRNAs as well as in positive and negative selection screens. We then examined the effects of 16,452 genetic perturbations on the abundance of eleven proteins representing a variety of cellular functions. We uncovered hundreds of regulatory relationships, including a novel link between the GAPDH isoenzymes Tdh1/2/3 and the Ras/PKA pathway. Many of the identified regulators are specific to one of the eleven proteins, but we also found genes that, upon perturbation, affected the abundance of most of the tested proteins. While the more specific regulators usually act transcriptionally, broad regulators often have roles in protein translation. Overall, our novel screening approach provides unprecedented insights into the components, scale and connectedness of the protein regulatory network.

Site-specific CRISPR-based mitochondrial DNA manipulation is limited by gRNA import.

Achieving CRISPR Cas9-based manipulation of mitochondrial DNA (mtDNA) has been a long-standing goal and would be of great relevance for disease modeling and for clinical applications. In this project, we aimed to deliver Cas9 into the mitochondria of human cells and analyzed Cas9-induced mtDNA cleavage and measured the resulting mtDNA depletion with multiplexed qPCR. In initial experiments, we found that measuring subtle effects on mtDNA copy numbers is challenging because of high biological variability, and detected no significant Cas9-caused mtDNA degradation. To overcome the challenge of being able to detect Cas9 activity on mtDNA, we delivered cytosine base editor Cas9-BE3 to mitochondria and measured its effect (C → T mutations) on mtDNA. Unlike regular Cas9-cutting, this leaves a permanent mark on mtDNA that can be detected with amplicon sequencing, even if the efficiency is low. We detected low levels of C → T mutations in cells that were exposed to mitochondrially targeted Cas9-BE3, but, surprisingly, these occurred regardless of whether a guide RNA (gRNA) specific to the targeted site, or non-targeting gRNA was used. This unspecific off-target activity shows that Cas9-BE3 can technically edit mtDNA, but also strongly indicates that gRNA import to mitochondria was not successful. Going forward mitochondria-targeted Cas9 base editors will be a useful tool for validating successful gRNA delivery to mitochondria without the ambiguity of approaches that rely on quantifying mtDNA copy numbers.

Characterization of Impurities in Therapeutic RNAs at the Single Nucleotide Level.

The chemistry of guide RNA (gRNA) affects the performance of the clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9 genome editing technique. However, the literature is very scarce about the study of gRNA degradation and in particular at the single nucleotide level. In this work, we developed a workflow to characterize the impurities of large RNAs at the single nucleotide level and identified the residues prone to degradation. Our strategy involves (i) the reduction of RNA length, (ii) a chromatographic mode able to capture subtle changes in impurity polarity, and (iii) a streamlined data treatment. To illustrate the approach, stressed gRNA samples were analyzed by coupling an immobilized ribonuclease T1 cartridge to a hydrophilic interaction liquid chromatography (HILIC) column hyphenated with tandem mass spectrometry (MS/MS). Critical findings were made possible by the presented technology. In particular, the desulfurization of phosphorothioate (PS) linkages was the major degradation observed at the single nucleotide level while no change in purity profile could be observed when using conventional ion-pairing reversed-phase (IPRP) liquid chromatography. To our knowledge, this is the first time that several impurity types are screened for a large RNA molecule using an automated online digestion analysis approach.

Identification and Analysis of Small Molecule Inhibitors of CRISPR-Cas9 in Human Cells.

Genome editing tools based on CRISPR-Cas systems can repair genetic mutations in situ; however, off-target effects and DNA damage lesions that result from genome editing remain major roadblocks to its full clinical implementation. Protein and chemical inhibitors of CRISPR-Cas systems may reduce off-target effects and DNA damage. Here we describe the identification of several lead chemical inhibitors that could specifically inhibit the activity of Streptococcus pyogenes Cas9 (SpCas9). In addition, we obtained derivatives of lead inhibitors that could penetrate the cell membrane and inhibit SpCas9 in cellulo. Two of these compounds, SP2 and SP24, were able to improve the specificity of SpCas9 in cellulo at low-micromolar concentration. Furthermore, microscale thermophoresis (MST) assays showed that SP24 might inhibit SpCas9 activity by interacting with both the SpCas9 protein and the SpCas9-gRNA ribonucleoprotein complex. Taken together, SP24 is a novel chemical inhibitor of SpCas9 which has the potential to enhance therapies that utilize SpCas9.

Host Cell Transcriptional Tuning with CRISPR/dCas9 to Mitigate the Effects of Toxin Exposure.

Anthrax infection is caused byBacillus anthracis, a bacterium that once established within the host releases lethal toxin (LeTx). Anthrax LeTx is internalized by the capillary morphogenesis protein 2/anthrax toxin receptor 2 (CMG2/ANTXR2) cell surface receptor on mammalian cells. Once inside the cell, LeTx cleaves mitogen-activated protein kinases (MAPKs), ultimately leading to cell death. Previous reports have shown that decreased expression of ANTXR2 reduces cell susceptibility to LeTx. By ablating the ANTXR2 gene in cells in vitro, we observed complete resistance to LeTx-induced cell death. Here, we directed CRISPR/dCas9-based tools to the ANTXR2 promoter to modulate ANTXR2 expression without altering the underlying gene sequence in human cell lines that express the receptor at high levels. We hypothesized that downregulating the expression of the ANTXR2 gene at the genomic level would mitigate the impact of toxin exposure. In one epigenetic editing approach, we employed the fusion of DNMT3A DNA methyltransferase and dCas9 (dCas9-DNMT3A) to methylate CpGs within the CpG island of the ANTXR2 promoter and found this repressed ANTXR2 gene expression resulting in significant resistance to LeTx-induced cell death. Furthermore, by multiplexing gRNAs to direct dCas9-DNMT3A to multiple sites in the ANTXR2 promoter, we applied a broader distribution of CpG methylation along the gene promoter resulting in enhanced repression and resistance to LeTx. In parallel, we directed the dCas9-KRAB-MeCP2 transcriptional repressor to the ANTXR2 promoter to quickly and robustly repress ANTXR2 expression. With this approach, in as little as two weeks, we created resistance to LeTx at a similar level to ANTXR2 gene-ablated cells. Overall, we present a transcriptional tuning approach to inhibit the effects of LeTx and provide a framework to repress toxin-binding cell surface receptors.

Multiplexed measurement of cell type-specific calcium kinetics using high-content image analysis combined with targeted gene disruption.

Kinetic analysis of intracellular calcium (Ca2+) in cardiomyocytes is commonly used to determine the pathogenicity of genetic mutations identified in patients with dilated cardiomyopathy (DCM). Conventional methods for measuring Ca2+ kinetics target whole-well cultured cardiomyocytes and therefore lack information concerning individual cells. Results are also affected by heterogeneity in cell populations. Here, we developed an analytical method using CRISPR/Cas9 genome editing combined with high-content image analysis (HCIA) that links cell-by-cell Ca2+ kinetics and immunofluorescence images in thousands of cardiomyocytes at a time. After transfecting cultured mouse cardiomyocytes that constitutively express Cas9 with gRNAs, we detected a prolonged action potential duration specifically in Serca2a-depleted ventricular cardiomyocytes in mixed culture. To determine the phenotypic effect of a frameshift mutation in PKD1 in a patient with DCM, we introduced the mutation into Cas9-expressing cardiomyocytes by gRNA transfection and found that it decreases the expression of PKD1-encoded PC1 protein that co-localizes specifically with Serca2a and L-type voltage-gated calcium channels. We also detected the suppression of Ca2+ amplitude in ventricular cardiomyocytes with decreased PC1 expression in mixed culture. Our HCIA method provides comprehensive kinetic and static information on individual cardiomyocytes and allows the pathogenicity of mutations to be determined rapidly.

Frankenstein Cas9: engineering improved gene editing systems.

The discovery of CRISPR-Cas9 and its widespread use has revolutionised and propelled research in biological sciences. Although the ability to target Cas9's nuclease activity to specific sites via an easily designed guide RNA (gRNA) has made it an adaptable gene editing system, it has many characteristics that could be improved for use in biotechnology. Cas9 exhibits significant off-target activity and low on-target nuclease activity in certain contexts. Scientists have undertaken ambitious protein engineering campaigns to bypass these limitations, producing several promising variants of Cas9. Cas9 variants with improved and alternative activities provide exciting new tools to expand the scope and fidelity of future CRISPR applications.

G-quadruplex-guided RNA engineering to modulate CRISPR-based genomic regulation.

It is important to develop small moelcule-based methods to modulate gene editing and expression in human cells. The roles of the G-quadruplex (G4) in biological systems have been widely studied. Here, G4-guided RNA engineering is performed to generate guide RNA with G4-forming units (G4-gRNA). We further demonstrate that chemical targeting of G4-gRNAs holds promise as a general approach for modulating gene editing and expression in human cells. The rich structural diversity of RNAs offers a reservoir of targets for small molecules to bind, thus creating the potential to modulate RNA biology.

Specificity of oligonucleotide gene therapy (OGT) agents.

Oligonucleotide gene therapy (OGT) agents (e. g. antisense, deoxyribozymes, siRNA and CRISPR/Cas) are promising therapeutic tools. Despite extensive efforts, only few OGT drugs have been approved for clinical use. Besides the problem of efficient delivery to targeted cells, hybridization specificity is a potential limitation of OGT agents. To ensure tight binding, a typical OGT agent hybridizes to the stretch of 15-25 nucleotides of a unique targeted sequence. However, hybrids of such lengths tolerate one or more mismatches under physiological conditions, the problem known as the affinity/specificity dilemma. Here, we assess the scale of this problem by analyzing OGT hybridization-dependent off-target effects (HD OTE) in vitro, in animal models and clinical studies. All OGT agents except deoxyribozymes exhibit HD OTE in vitro, with most thorough evidence of poor specificity reported for siRNA and CRISPR/Cas9. Notably, siRNA suppress non-targeted genes due to (1) the partial complementarity to mRNA 3'-untranslated regions (3'-UTR), and (2) the antisense activity of the sense strand. CRISPR/Cas9 system can cause hundreds of non-intended dsDNA breaks due to low specificity of the guide RNA, which can limit therapeutic applications of CRISPR/Cas9 by ex-vivo formats. Contribution of this effects to the observed in vivo toxicity of OGT agents is unclear and requires further investigation. Locked or peptide nucleic acids improve OGT nuclease resistance but not specificity. Approaches that use RNA marker dependent (conditional) activation of OGT agents may improve specificity but require additional validation in cell culture and in vivo.

Small Molecules for Enhancing the Precision and Safety of Genome Editing.

Clustered regularly interspaced short palindromic repeats (CRISPR)-based genome-editing technologies have revolutionized biology, biotechnology, and medicine, and have spurred the development of new therapeutic modalities. However, there remain several barriers to the safe use of CRISPR technologies, such as unintended off-target DNA cleavages. Small molecules are important resources to solve these problems, given their facile delivery and fast action to enable temporal control of the CRISPR systems. Here, we provide a comprehensive overview of small molecules that can precisely modulate CRISPR-associated (Cas) nucleases and guide RNAs (gRNAs). We also discuss the small-molecule control of emerging genome editors (e.g., base editors) and anti-CRISPR proteins. These molecules could be used for the precise investigation of biological systems and the development of safer therapeutic modalities.

Enzyme-free targeted DNA demethylation using CRISPR-dCas9-based steric hindrance to identify DNA methylation marks causal to altered gene expression.

DNA methylation involves the enzymatic addition of a methyl group primarily to cytosine residues in DNA. This protocol describes how to produce complete and minimally confounded DNA demethylation of specific sites in the genome of cultured cells by clustered regularly interspaced short palindromic repeats (CRISPR)-dCas9 and without the involvement of an epigenetic-modifying enzyme, the purpose of which is the evaluation of the functional (i.e., gene expression or phenotypic) consequences of DNA demethylation of specific sites that have been previously implicated in particular pathological or physiological contexts. This protocol maximizes the ability of the easily reprogrammable CRISPR-dCas9 system to assess the impact of DNA methylation from a causal rather than correlational perspective: alternative protocols for CRISPR-dCas9-based site-specific DNA methylation or demethylation rely on the recruitment of epigenetic enzymes that exhibit additional nonspecific activities at both the targeted site and throughout the genome, confounding conclusions of causality of DNA methylation. Inhibition or loss of DNA methylation is accomplished by three consecutive lentiviral transductions. The first two lentiviruses establish stable expression of dCas9 and a guide RNA, which will physically obstruct either maintenance or de novo DNA methyltransferase activity at the guide RNA target site. A third lentivirus introduces Cre recombinase to delete the dCas9 transgene, which leads to loss of dCas9 from the target site, allowing transcription factors and/or the transcription machinery to interact with the demethylated target site. This protocol requires 3-8 months to complete owing to prolonged cell passaging times, but there is little hands-on time, and no specific skills beyond basic molecular biology techniques are necessary.