Multiplex gene precise editing and large DNA fragment deletion by the CRISPR-Cas9-TRAMA system in edible mushroom Cordyceps militaris

The medicinal mushroom Cordyceps militaris contains abundant valuable bioactive ingredients that have attracted a great deal of attention in the pharmaceutical and cosmetic industries. However, the development of this valuable mushroom faces the obstacle of lacking powerful genomic engineering tools. Here, by excavating the endogenous tRNA-processed element, introducing the extrachromosomal plasmid and alongside with homologous template, we develop a marker-free CRISPR-Cas9-TRAMA genomic editing system to achieve the multiplex gene precise editing and large synthetic cluster deletion in C. militaris. We further operated editing in the synthetases of cordycepin and ergothioneine to demonstrate the application of Cas9-TRAMA system in protein modification, promoter strength evaluation and 10 kb metabolic synthetic cluster deletion. The Cas9-TRAMA system provides a scalable method for excavating the valuable metabolic resource of medicinal mushrooms and constructing a mystical cellular pathway to elucidate the complex cell behaviours of the edible mushroom.     

A new family of CRISPR‐type V nucleases with C‐rich PAM recognition

Most CRISPR‐type V nucleases are stimulated to cleave double‐stranded (ds) DNA targets by a T‐rich PAM, which restricts their targeting range. Here, we identify and characterize a new family of type V RNA‐guided nuclease, Cas12l, that exclusively recognizes a C‐rich (5''‐CCY‐3′) PAM. The organization of genes within its CRISPR locus is similar to type II‐B CRISPR‐Cas9 systems, but both sequence analysis and functional studies establish it as a new family of type V effector. Biochemical experiments show that Cas12l nucleases function optimally between 37 and 52°C, depending on the ortholog, and preferentially cut supercoiled DNA. Like other type V nucleases, it exhibits collateral nonspecific ssDNA and ssRNA cleavage activity that is triggered by ssDNA or dsDNA target recognition. Finally, we show that one family member, Asp2Cas12l, functions in a heterologous cellular environment, altogether, suggesting that this new group of CRISPR‐associated nucleases may be harnessed as genome editing reagents.     

Expanding the plant genome editing toolbox with recently developed CRISPR–Cas systems

Since its first appearance, CRISPR–Cas9 has been developed extensively as a programmable genome-editing tool, opening a new era in plant genome engineering. However, CRISPR–Cas9 still has some drawbacks, such as limitations of the protospacer-adjacent motif (PAM) sequence, target specificity, and the large size of the cas9 gene. To combat invading bacterial phages and plasmid DNAs, bacteria and archaea have diverse and unexplored CRISPR–Cas systems, which have the potential to be developed as a useful genome editing tools. Recently, discovery and characterization of additional CRISPR–Cas systems have been reported. Among them, several CRISPR–Cas systems have been applied successfully to plant and human genome editing. For example, several groups have achieved genome editing using CRISPR–Cas type I-D and type I-E systems, which had never been applied for genome editing previously. In addition to higher specificity and recognition of different PAM sequences, recently developed CRISPR–Cas systems often provide unique characteristics that differ from well-known Cas proteins such as Cas9 and Cas12a. For example, type I CRISPR–Cas10 induces small indels and bi-directional long-range deletions ranging up to 7.2 kb in tomatoes (Solanum lycopersicum L.). Type IV CRISPR–Cas13 targets RNA, not double-strand DNA, enabling highly specific knockdown of target genes. In this article, we review the development of CRISPR–Cas systems, focusing especially on their application to plant genome engineering. Recent CRISPR–Cas tools are helping expand our plant genome engineering toolbox.

Recently discovered and characterized clustered regularly interspaced short palindromic repeats-CRISPR associated (CRISPR–Cas) systems allow additional applications to plant genome editing.     

Between a rock and a hard place: Farmers’ perspectives on gene editing in livestock agriculture in Bavaria

Farmers’ opinions and concerns are rarely considered in public debates about the use of gene‐editing technologies to modify farm animals. Subject Categories: Synthetic Biology & Biotechnology, S&S: Economics & Business

CRISPR‐Cas9, a new method for precisely modifying DNA, has received significant public attention in recent years. Heralded as a breakthrough technology to genetically modify organisms with hitherto unknown ease, precision and at low cost, CRISPR‐Cas9 has reignited controversies in science and society about the kind of genetic modifications that can and should be achieved in animals, plants and humans. Historically, such public debates have been particularly heated in two areas: human germline modification and agricultural applications. This article focuses on the latter and explores how small‐ and medium‐scale farmers evaluate the possibility and the potential benefits of gene editing livestock. This article importantly adds their voices to the discussion – voices that are surprisingly often unheard or ignored in public debates about using genetic technologies in agriculture.

… CRISPR‐Cas9 has reignited controversies in science and society about the kind of genetic modifications that can and should be achieved in animals, plants and humans.

     

Unexpected gene activation following CRISPR‐Cas9‐mediated genome editing

The discovery of the Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) and its development as a genome editing tool has revolutionized the field of molecular biology. In the DNA damage field, CRISPR has brought an alternative to induce endogenous double‐strand breaks (DSBs) at desired genomic locations and study the DNA damage response and its consequences. Many systems for sgRNA delivery have been reported in order to efficiently generate this DSB, including lentiviral vectors. However, some of the consequences of these systems are not yet well understood. Here, we report that lentiviral‐based sgRNA vectors can integrate into the endogenous genomic target location, leading to undesired activation of the target gene. By generating a DSB in the regulatory region of the ABCB1 gene using a lentiviral sgRNA vector, we can induce the formation of Taxol‐resistant colonies. We show that these colonies upregulate ABCB1 via integration of the EEF1A1 and the U6 promoters from the sgRNA vector. We believe that this is an unreported CRISPR/Cas9 on‐target effect that researchers need to be aware of when using lentiviral vectors for genome editing.     

Chemogenetic profiling reveals PP2A‐independent cytotoxicity of proposed PP2A activators iHAP1 and DT‐061

Protein phosphatase 2A (PP2A) is an abundant phosphoprotein phosphatase that acts as a tumor suppressor. For this reason, compounds able to activate PP2A are attractive anticancer agents. The compounds iHAP1 and DT‐061 have recently been reported to selectively stabilize specific PP2A‐B56 complexes to mediate cell killing. We were unable to detect direct effects of iHAP1 and DT‐061 on PP2A‐B56 activity in biochemical assays and composition of holoenzymes. Therefore, we undertook genome‐wide CRISPR‐Cas9 synthetic lethality screens to uncover biological pathways affected by these compounds. We found that knockout of mitotic regulators is synthetic lethal with iHAP1 while knockout of endoplasmic reticulum (ER) and Golgi components is synthetic lethal with DT‐061. Indeed we showed that iHAP1 directly blocks microtubule assembly both in vitro and in vivo and thus acts as a microtubule poison. In contrast, DT‐061 disrupts both the Golgi apparatus and the ER and lipid synthesis associated with these structures. Our work provides insight into the biological pathways perturbed by iHAP1 and DT‐061 causing cellular toxicity and argues that these compounds cannot be used for dissecting PP2A‐B56 biology.     

CRISPR–Cas9-assisted recombineering in Lactobacillus reuteri

Clustered regularly interspaced palindromic repeats (CRISPRs) and the CRISPR-associated (Cas) nuclease protect bacteria and archeae from foreign DNA by site-specific cleavage of incoming DNA. Type-II CRISPR–Cas systems, such as the Streptococcus pyogenes CRISPR–Cas9 system, can be adapted such that Cas9 can be guided to a user-defined site in the chromosome to introduce double-stranded breaks. Here we have developed and optimized CRISPR–Cas9 function in the lactic acid bacterium Lactobacillus reuteri ATCC PTA 6475. We established proof-of-concept showing that CRISPR–Cas9 selection combined with single-stranded DNA (ssDNA) recombineering is a realistic approach to identify at high efficiencies edited cells in a lactic acid bacterium. We show for three independent targets that subtle changes in the bacterial genome can be recovered at efficiencies ranging from 90 to 100%. By combining CRISPR–Cas9 and recombineering, we successfully applied codon saturation mutagenesis in the L. reuteri chromosome. Also, CRISPR–Cas9 selection is critical to identify low-efficiency events such as oligonucleotide-mediated chromosome deletions. This also means that CRISPR–Cas9 selection will allow identification of recombinant cells in bacteria with low recombineering efficiencies, eliminating the need for ssDNA recombineering optimization procedures. We envision that CRISPR–Cas genome editing has the potential to change the landscape of genome editing in lactic acid bacteria, and other Gram-positive bacteria.     

An efficient and precise method for generating knockout cell lines based on CRISPR‐Cas9 system

Although the efficiency and versatility of CRISPR‐Cas9 system has been greatly improved over conventional genome editing methods such as zinc finger or TALEN, it is still time‐consuming and labor‐intensive for screening knockout/knock‐in cell clones due to differences of the targeted location or efficacies of guide RNAs (gRNAs). Here, we adapted a targeted knock‐in strategy with CRISPR‐Cas9 system and characterized the efficiency for generating single or double knockout cell lines. Specifically, a homology‐arm based donor cassette consisting of genes encoding a fluorescence protein and antibiotic selection marker driven by a constitutive promoter was co‐transfected with a gRNA expressing unit. Based on FACS sorting and antibiotic drug selection, positive cell clones were confirmed by genotyping and at the protein expression level. The results indicated that more than 70% of analyzed clones identified by cell sorting and selection were successfully targeted in both single and double knockout experiments. The procedure takes less than three weeks to obtain knockout cell lines. We believe that this methodology could be applicable and versatile in generating knockout cell clones with high efficiency in most cell lines.     

Customized optical mapping by CRISPR–Cas9 mediated DNA labeling with multiple sgRNAs

Whole-genome mapping technologies have been developed as a complementary tool to provide scaffolds for genome assembly and structural variation analysis (1,2). We recently introduced a novel DNA labeling strategy based on a CRISPR–Cas9 genome editing system, which can target any 20bp sequences. The labeling strategy is specifically useful in targeting repetitive sequences, and sequences not accessible to other labeling methods. In this report, we present customized mapping strategies that extend the applications of CRISPR–Cas9 DNA labeling. We first design a CRISPR–Cas9 labeling strategy to interrogate and differentiate the single allele differences in NGG protospacer adjacent motifs (PAM sequence). Combined with sequence motif labeling, we can pinpoint the single-base differences in highly conserved sequences. In the second strategy, we design mapping patterns across a genome by selecting sets of specific single-guide RNAs (sgRNAs) for labeling multiple loci of a genomic region or a whole genome. By developing and optimizing a single tube synthesis of multiple sgRNAs, we demonstrate the utility of CRISPR–Cas9 mapping with 162 sgRNAs targeting the 2Mb Haemophilus influenzae chromosome. These CRISPR–Cas9 mapping approaches could be particularly useful for applications in defining long-distance haplotypes and pinpointing the breakpoints in large structural variants in complex genomes and microbial mixtures.     

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.     

Epigenetic features drastically impact CRISPR–Cas9 efficacy in plants

CRISPR–Cas9-mediated genome editing has been widely adopted for basic and applied biological research in eukaryotic systems. While many studies consider DNA sequences of CRISPR target sites as the primary determinant for CRISPR mutagenesis efficiency and mutation profiles, increasing evidence reveals the substantial role of chromatin context. Nonetheless, most prior studies are limited by the lack of sufficient epigenetic resources and/or by only transiently expressing CRISPR–Cas9 in a short time window. In this study, we leveraged the wealth of high-resolution epigenomic resources in Arabidopsis (Arabidopsis thaliana) to address the impact of chromatin features on CRISPR–Cas9 mutagenesis using stable transgenic plants. Our results indicated that DNA methylation and chromatin features could lead to substantial variations in mutagenesis efficiency by up to 250-fold. Low mutagenesis efficiencies were mostly associated with repressive heterochromatic features. This repressive effect appeared to persist through cell divisions but could be alleviated through substantial reduction of DNA methylation at CRISPR target sites. Moreover, specific chromatin features, such as H3K4me1, H3.3, and H3.1, appear to be associated with significant variation in CRISPR–Cas9 mutation profiles mediated by the non-homologous end joining repair pathway. Our findings provide strong evidence that specific chromatin features could have substantial and lasting impacts on both CRISPR–Cas9 mutagenesis efficiency and DNA double-strand break repair outcomes.

Epigenetic features substantially influence genome editing efficiency and DNA repair outcomes.     

High‐efficiency delivery of CRISPR‐Cas9 by engineered probiotics enables precise microbiome editing

Antibiotic resistance threatens our ability to treat infectious diseases, spurring interest in alternative antimicrobial technologies. The use of bacterial conjugation to deliver CRISPR‐cas systems programmed to precisely eliminate antibiotic‐resistant bacteria represents a promising approach but requires high in situ DNA transfer rates. We have optimized the transfer efficiency of conjugative plasmid TP114 using accelerated laboratory evolution. We hence generated a potent conjugative delivery vehicle for CRISPR‐cas9 that can eliminate > 99.9% of targeted antibiotic‐resistant Escherichia coli in the mouse gut microbiota using a single dose. We then applied this system to a Citrobacter rodentium infection model, achieving full clearance within four consecutive days of treatment.     

Efficient genome editing in Caenorhabditis elegans by CRISPR-targeted homologous recombination

Cas9 is an RNA-guided double-stranded DNA nuclease that participates in clustered regularly interspaced short palindromic repeats (CRISPR)-mediated adaptive immunity in prokaryotes. CRISPR–Cas9 has recently been used to generate insertion and deletion mutations in Caenorhabditis elegans, but not to create tailored changes (knock-ins). We show that the CRISPR–CRISPR-associated (Cas) system can be adapted for efficient and precise editing of the C. elegans genome. The targeted double-strand breaks generated by CRISPR are substrates for transgene-instructed gene conversion. This allows customized changes in the C. elegans genome by homologous recombination: sequences contained in the repair template (the transgene) are copied by gene conversion into the genome. The possibility to edit the C. elegans genome at selected locations will facilitate the systematic study of gene function in this widely used model organism.     

Discovery of potent and versatile CRISPR–Cas9 inhibitors engineered for chemically controllable genome editing

Anti-CRISPR (Acr) proteins are encoded by many mobile genetic elements (MGEs) such as phages and plasmids to combat CRISPR–Cas adaptive immune systems employed by prokaryotes, which provide powerful tools for CRISPR–Cas-based applications. Here, we discovered nine distinct type II-A anti-CRISPR (AcrIIA24–32) families from Streptococcus MGEs and found that most Acrs can potently inhibit type II-A Cas9 orthologs from Streptococcus (SpyCas9, St1Cas9 or St3Cas9) in bacterial and human cells. Among these Acrs, AcrIIA26, AcrIIA27, AcrIIA30 and AcrIIA31 are able to block Cas9 binding to DNA, while AcrIIA24 abrogates DNA cleavage by Cas9. Notably, AcrIIA25.1 and AcrIIA32.1 can inhibit both DNA binding and DNA cleavage activities of SpyCas9, exhibiting unique anti-CRISPR characteristics. Importantly, we developed several chemically inducible anti-CRISPR variants based on AcrIIA25.1 and AcrIIA32.1 by comprising hybrids of Acr protein and the 4-hydroxytamoxifen-responsive intein, which enabled post-translational control of CRISPR–Cas9-mediated genome editing in human cells. Taken together, our work expands the diversity of type II-A anti-CRISPR families and the toolbox of Acr proteins for the chemically inducible control of Cas9-based applications.     

Alzheimer’s disease associated AKAP9 I2558M mutation alters posttranslational modification and interactome of tau and cellular functions in CRISPR‐edited human neuronal cells

Alzheimer''s disease (AD) is a pervasive neurodegeneration disease with high heritability. In this study, we employed CRISPR‐Cas9‐engineered technology to investigate the effects of a rare mutation (rs144662445) in the A kinase anchoring protein 9 (AKAP9) gene, which is associated with AD in African Americans (AA), on tau pathology and the tau interactome in SH‐SY5Y P301L neuron‐like cells. The mutation significantly increased the level of phosphorylated tau, specifically at the site Ser396/Ser404. Moreover, analyses of the tau interactome measured by affinity purification‐mass spectrometry revealed that differentially expressed tau‐interacting proteins in AKAP9 mutant cells were associated with RNA translation, RNA localization and oxidative activity, recapitulating the tau interactome signature previously reported with human AD brain samples. Importantly, these results were further validated by functional studies showing a significant reduction in protein synthesis activity and excessive oxidative stress in AKAP9 mutant compared with wild type cells in a tau‐dependent manner, which are mirrored with pathological phenotype frequently seen in AD. Our results demonstrated specific effects of rs14462445 on mis‐processing of tau and suggest a potential role of AKAP9 in AD pathogenesis.     

Systematic in vitro specificity profiling reveals nicking defects in natural and engineered CRISPR–Cas9 variants

Cas9 is an RNA-guided endonuclease in the bacterial CRISPR–Cas immune system and a popular tool for genome editing. The commonly used Streptococcus pyogenes Cas9 (SpCas9) is relatively non-specific and prone to off-target genome editing. Other Cas9 orthologs and engineered variants of SpCas9 have been reported to be more specific. However, previous studies have focused on specificity of double-strand break (DSB) or indel formation, potentially overlooking alternative cleavage activities of these Cas9 variants. In this study, we employed in vitro cleavage assays of target libraries coupled with high-throughput sequencing to systematically compare cleavage activities and specificities of two natural Cas9 variants (SpCas9 and Staphylococcus aureus Cas9) and three engineered SpCas9 variants (SpCas9 HF1, HypaCas9 and HiFi Cas9). We observed that all Cas9s tested could cleave target sequences with up to five mismatches. However, the rate of cleavage of both on-target and off-target sequences varied based on target sequence and Cas9 variant. In addition, SaCas9 and engineered SpCas9 variants nick targets with multiple mismatches but have a defect in generating a DSB, while SpCas9 creates DSBs at these targets. Overall, these differences in cleavage rates and DSB formation may contribute to varied specificities observed in genome editing studies.     

Structural basis for inhibition of the type I-F CRISPR–Cas surveillance complex by AcrIF4, AcrIF7 and AcrIF14

CRISPR–Cas systems are adaptive immune systems in bacteria and archaea to defend against mobile genetic elements (MGEs) and have been repurposed as genome editing tools. Anti-CRISPR (Acr) proteins are produced by MGEs to counteract CRISPR–Cas systems and can be used to regulate genome editing by CRISPR techniques. Here, we report the cryo-EM structures of three type I-F Acr proteins, AcrIF4, AcrIF7 and AcrIF14, bound to the type I-F CRISPR–Cas surveillance complex (the Csy complex) from Pseudomonas aeruginosa. AcrIF4 binds to an unprecedented site on the C-terminal helical bundle of Cas8f subunit, precluding conformational changes required for activation of the Csy complex. AcrIF7 mimics the PAM duplex of target DNA and is bound to the N-terminal DNA vise of Cas8f. Two copies of AcrIF14 bind to the thumb domains of Cas7.4f and Cas7.6f, preventing hybridization between target DNA and the crRNA. Our results reveal structural detail of three AcrIF proteins, each binding to a different site on the Csy complex for inhibiting degradation of MGEs.