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Exploration of Hygromycin B Biosynthesis Utilizing CRISPR-Cas9-Associated Base Editing.
Hygromycin B is an aminoglycoside antibiotic widely used in industry and biological research. However, most of its biosynthetic pathway has not been completely identified due to the immense difficulty in genetic manipulation of the producing strain. To address this problem, we developed an efficient system that combines clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9-associated base editing and site-specific recombination instead of conventional double-crossover-based homologous recombination. This strategy was successfully applied to the in vivo inactivation of five candidate genes involved in the biosynthesis of hygromycin B by generating stop codons or mutating conserved residues within the encoding region. The results revealed that HygJ, HygL, and HygD are responsible for successive dehydrogenation, transamination, and transglycosylation of nucleoside diphosphate (NDP)-heptose. Notably, HygY acts as an unusual radical S-adenosylmethionine (SAM)-dependent epimerase for hydroxyl carbons, and HygM serves as a versatile methyltransferase in multiple parallel metabolic networks. Based on in vivo and in vitro evidence, the biosynthetic pathway for hygromycin B is proposed.
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The pAblo·pCasso self-curing vector toolset for unconstrained cytidine and adenine base-editing in Gram-negative bacteria.
A synthetic biology toolkit, exploiting clustered regularly interspaced short palindromic repeats (CRISPR) and modified CRISPR-associated protein (Cas) base-editors, was developed for genome engineering in Gram-negative bacteria. Both a cytidine base-editor (CBE) and an adenine base-editor (ABE) have been optimized for precise single-nucleotide modification of plasmid and genome targets. CBE comprises a cytidine deaminase conjugated to a Cas9 nickase from Streptococcus pyogenes (SpnCas9), resulting in C→T (or G→A) substitutions. Conversely, ABE consists of an adenine deaminase fused to SpnCas9 for A→G (or T→C) editing. Several nucleotide substitutions were achieved using these plasmid-borne base-editing systems and a novel protospacer adjacent motif (PAM)-relaxed SpnCas9 (SpRY) variant. Base-editing was validated in Pseudomonas putida and other Gram-negative bacteria by inserting premature STOP codons into target genes, thereby inactivating both fluorescent proteins and metabolic (antibiotic-resistance) functions. The functional knockouts obtained by engineering STOP codons via CBE were reverted to the wild-type genotype using ABE. Additionally, a series of induction-responsive vectors was developed to facilitate the curing of the base-editing platform in a single cultivation step, simplifying complex strain engineering programs without relying on homologous recombination and yielding plasmid-free, modified bacterial cells.
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Light-induced expression of gRNA allows for optogenetic gene editing of T lymphocytes in vivo.
There is currently a lack of tools capable of perturbing genes in both a precise and a spatiotemporal fashion. The flexibility of CRISPR (clustered regularly interspaced short palindromic repeats), coupled with light's unparalleled spatiotemporal resolution deliverable from a controllable source, makes optogenetic CRISPR a well-suited solution for precise spatiotemporal gene perturbations. Here, we present a new optogenetic CRISPR tool (Blue Light-inducible Universal VPR-Improved Production of RGRs, BLU-VIPR) that diverges from prevailing split-Cas design strategies and instead focuses on optogenetic regulation of guide RNA (gRNA) production. We engineered BLU-VIPR around a new potent blue-light activated transcription factor (VPR-EL222) and ribozyme-flanked gRNA. The BLU-VIPR design is genetically encoded and ensures precise excision of multiple gRNAs from a single messenger RNA transcript. This simplified spatiotemporal gene perturbation and allowed for several types of optogenetic CRISPR, including indels, CRISPRa, and base editing. BLU-VIPR also worked in vivo with cells previously intractable to optogenetic gene editing, achieving optogenetic gene editing in T lymphocytes in vivo.
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Programmable RNA base editing via targeted modifications.
Clustered regularly interspaced short palindromic repeats (CRISPR)-based genome editors are powerful tools in biology and hold great promise for the treatment of human diseases. Advanced DNA base editing tools, such as cytosine base editor and adenine base editor, have been developed to correct permanent mistakes in genetic material. However, undesired off-target edits would also be permanent, which poses a considerable risk for therapeutics. Alternatively, base editing at the RNA level is capable of correcting disease-causing mutations but does not lead to lasting genotoxic effects. RNA base editors offer temporary and reversible therapies and have been catching on in recent years. Here, we summarize some emerging RNA editors based on A-to-inosine, C-to-U and U-to-pseudouridine changes. We review the programmable RNA-targeting systems as well as modification enzyme-based effector proteins and highlight recent technological breakthroughs. Finally, we compare editing tools, discuss limitations and opportunities, and provide insights for the future directions of RNA base editing.
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Cationic Polymer-Mediated CRISPR/Cas9 Plasmid Delivery for Genome Editing.
Delivery of CRISPR (clustered regularly interspaced short palindromic repeats)/CRISPR-associated protein-9 (Cas9) represents a major hurdle for successful clinical translation of genome editing tools. Owing to the large size of plasmids that encode Cas9 and single-guide RNA (sgRNA), genome editing efficiency mediated by current delivery carriers is still unsatisfactory to meet the requirement for its real applications. Herein, cationic polymer polyethyleneimine-β-cyclodextrin (PC), known to be efficient for small plasmid transfection, is reported to likewise mediate efficient delivery of plasmid encoding Cas9 and sgRNA. Whereas PC can condense and encapsulate large plasmids at high N/P ratio, the delivery of plasmid results in efficient editing at two genome loci, namely, hemoglobin subunit beta (19.1%) and rhomboid 5 homolog 1 (RHBDF1) (7.0%). Sanger sequencing further confirms the successful genome editing at these loci. This study defines a new strategy for the delivery of the large plasmid encoding Cas9/sgRNA for efficient genome editing.
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In Vivo Rapid Investigation of CRISPR-Based Base Editing Components in Escherichia coli (IRI-CCE): A Platform for Evaluating Base Editing Tools and Their Components.
Rapid assessment of clustered regularly interspaced short palindromic repeats/CRISPR-associated protein (CRISPR/Cas)-based genome editing (GE) tools and their components is a critical aspect for successful GE applications in different organisms. In many bacteria, double-strand breaks (DSBs) generated by CRISPR/Cas tool generally cause cell death due to the lack of an efficient nonhomologous end-joining pathway and restricts its use. CRISPR-based DSB-free base editors (BEs) have been applied for precise nucleotide (nt) editing in bacteria, which does not need to make DSBs. However, optimization of newer BE tools in bacteria is challenging owing to the toxic effects of BE reagents expressed using strong promoters. Improved variants of two main BEs, cytidine base editor (CBE) and adenine base editor (ABE), capable of converting C to T and A to G, respectively, have been recently developed but yet to be tested for editing characteristics in bacteria. Here, we report a platform for in vivo rapid investigation of CRISPR-BE components in Escherichia coli (IRI-CCE) comprising a combination of promoters and terminators enabling the expression of nCas9-based BE and sgRNA to nontoxic levels, eventually leading to successful base editing. We demonstrate the use of IRI-CCE to characterize different variants of CBEs (PmCDA1, evoCDA1, APOBEC3A) and ABEs (ABE8e, ABE9e) for bacteria, exhibiting that each independent BE has its specific editing pattern for a given target site depending on protospacer length. In summary, CRISPR-BE components expressed without lethal effects on cell survival in the IRI-CCE allow an analysis of various BE tools, including cloned biopart modules and sgRNAs.
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Deletion of FOXL2 by CRISPR promotes cell cycle G0/G1 restriction in KGN cells.
Forkhead box L2 (FOXL2), a member of the forkhead family of transcription factors, is important in eyelid and ovary differentiation. Although the function of FOXL2 in organogenesis has been investigated, the detailed mechanisms by which FOXL2 mediates cellular process remain to be fully elucidated. Few FOXL2‑knockout cell lines have been reported, which has limited molecular mechanism investigations. CRISPR is a novel gene editing technique that has been widely used in human genetic diseases. In the present study, FOXL2 was disrupted using clustered regularly interspaced short palindromic repeats (CRISPR), and screening of a stable knockout cell line was performed in human ovarian granulosa KGN cells. Three sites (F404, F425 and F446) around the ATG start codon on the FOXL2 DNA sequence were constructed in a guide RNA lentivirus. Targeting F425 was most efficient, and western blot analysis and DNA sequencing of the resulting cells suggested that both FOXL2 alleles were fully disrupted. In addition, flow cytometry results indicated that the knockout of FOXL2 restricted cell cycle progression at the G0/G1 phase. In addition, the expression levels of cell cycle mediators cyclin D1 and cyclin‑dependent kinase 4 were reduced. These results confirmed that FOXL2 disruption in KGN cells is associated with the cell cycle attenuation.
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A systematic review of computational methods for designing efficient guides for CRISPR DNA base editor systems.
In only a few years, as a breakthrough technology, clustered regularly interspaced short palindromic repeats/CRISPR-associated protein (CRISPR/Cas) gene-editing systems have ushered in the era of genome engineering with a plethora of applications. One of the most promising CRISPR tools, so-called base editors, opened an exciting avenue for exploring new therapeutic approaches through controlled mutagenesis. However, the efficiency of a base editor guide varies depending on several biological determinants, such as chromatin accessibility, DNA repair proteins, transcriptional activity, factors related to local sequence context and so on. Thus, the success of genetic perturbation directed by CRISPR/Cas base-editing systems relies on an optimal single guide RNA (sgRNA) design, taking those determinants into account. Although there is 11 commonly used software to design guides specifically for base editors, only three of them investigated and implemented those biological determinants into their models. This review presents the key features, capabilities and limitations of all currently available software with a particular focus on predictive model-based algorithms. Here, we summarize existing software for sgRNA design and provide a base for improving the efficiency of existing available software suites for precise target base editing.
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Recent Advances in CRISPR/Cas Technologies for Biological Discovery, Therapeutics, and Diagnostics.
The clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein (Cas) system began as a tool for programmable genome editing. CRISPR/Cas technologies have evolved into a versatile platform for functional genomic screening, epigenome editing, therapeutic target discovery, and highly sensitive molecular diagnostics. New effectors and engineered variants continue to push these applications into personalized medicine and point-of-care testing. Here, this review highlights recent advances in therapeutic target discovery and therapeutic and molecular diagnostic development utilizing CRISPR/Cas technologies. We discuss how CRISPR interference (CRISPRi), CRISPR activation (CRISPRa), base editing, and prime editing have improved our understanding of disease mechanisms, while creating new opportunities for therapeutic intervention. Current applications in cancer immunotherapy, infectious disease, and neurological disorders are also discussed. In diagnostics, CRISPR-based platforms enable sensitive detection of infectious pathogens, cancer biomarkers, and genetic disorders using programmable nuclease activity in both laboratory and point-of-care settings. Collectively, these advances continue to expand the role of CRISPR/Cas technology across biological discovery, disease diagnostics, and therapeutic development, driving progress in precision medicine.
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Efficient gene editing in adult mouse livers via adenoviral delivery of CRISPR/Cas9.
We developed an adenovirus-based CRISPR/Cas9 system for gene editing in vivo. In the liver, we demonstrated that the system could reach the level of tissue-specific gene knockout, resulting in phenotypic changes. Given the wide spectrum of cell types susceptible to adenoviral infection, and the fact that adenoviral genome rarely integrates into its host cell genome, we believe the adenovirus-based CRISPR/Cas9 system will find applications in a variety of experimental settings.
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