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Found: 26 116
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Large Donor CRISPR for Whole-Coding Sequence Replacement of Cell Adhesion Molecule LRRTM2.
The cell adhesion molecule leucine-rich repeat transmembrane neuronal protein 2 (LRRTM2) is crucial for synapse development and function. However, our understanding of its endogenous trafficking has been limited due to difficulties in manipulating its coding sequence (CDS) using standard genome editing techniques. Instead, we replaced the entire LRRTM2 CDS by adapting a two-guide CRISPR knock-in method, enabling complete control of LRRTM2. In primary rat hippocampal cultures dissociated from embryos of both sexes, N-terminally tagged, endogenous LRRTM2 was found in 80% of synapses, and synaptic LRRTM2 content correlated with PSD-95 and AMPAR levels. LRRTM2 was also enriched with AMPARs outside synapses, demonstrating the sensitivity of this method to detect relevant new biology. Finally, we leveraged total genomic control to increase the synaptic levels of LRRTM2 via simultaneous mutation of its C-terminal domain, which did not correspondingly increase AMPAR enrichment. The coding region of thousands of genes span lengths suitable for whole-CDS replacement, suggesting this simple approach will enable straightforward structure-function analysis in neurons.
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Gene-edited MLE-15 Cells as a Model for the Hermansky-Pudlak Syndromes.
Defining the mechanisms of cellular pathogenesis in rare lung diseases such as Hermansky-Pudlak syndrome (HPS) is often complicated by loss of the differentiated phenotype of cultured primary alveolar type 2 (AT2) cells, as well as by a lack of durable cell lines that are faithful to both AT2-cell and rare disease phenotypes. We used CRISPR/Cas9 gene editing to generate a series of HPS-specific mutations in the MLE-15 cell line. The resulting MLE-15/HPS cell lines exhibit preservation of AT2 cellular functions, including formation of lamellar body-like organelles, complete processing of surfactant protein B, and known features of HPS specific to each trafficking complex, including loss of protein targeting to lamellar bodies. MLE-15/HPS1 and MLE-15/HPS2 (with a mutation in Ap3β1) express increased macrophage chemotactic protein-1, a well-described mediator of alveolitis in patients with HPS and in mouse models. We show that MLE-15/HPS9 and pallid AT2 cells (with a mutation in Bloc1s6) also express increased macrophage chemotactic protein-1, suggesting that mice and humans with BLOC-1 mutations may also be susceptible to alveolitis. In addition to providing a flexible platform to examine the role of HPS-specific mutations in trafficking AT2 cells, MLE-15/HPS cell lines provide a durable resource for high-throughput screening and studies of cellular pathophysiology that are likely to accelerate progress toward developing novel therapies for this rare lung disease.
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Precise virulence inactivation using a CRISPR-associated transposase for combating Enterobacteriaceae gut pathogens.
Targeted gene manipulation in a complex microbial community is an enabling technology for precise microbiome editing. Here we introduce BACTRINS, an in situ microbiome engineering platform designed for efficient and precise genomic insertion of a desired payload and simultaneous knockout of target genes. This system leverages conjugation-mediated delivery of CRISPR-associated transposases to achieve RNA-guided genomic integration, allowing precise insertion of a therapeutic payload while neutralizing pathogen virulence without causing cell death. When applied against an Enterobacteriaceae Shiga toxin-producing pathogen in the gut, this system delivers a CRISPR-associated transposase by bacterial conjugation for site-specific inactivation of the Shiga toxin gene and integration of a nanobody therapeutic payload to disrupt pathogen attachment. A single dose of this therapy results in high-efficiency Shiga gene inactivation and improved survival in a murine infection model of Shiga-producing pathogen. This work establishes a new type of live bacterial therapeutic capable of reducing gut infections by transforming toxigenic pathogens into commensal protectors.
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Filamentation and restoration of normal growth in Escherichia coli using a combined CRISPRi sgRNA/antisense RNA approach.
CRISPR interference (CRISPRi) using dCas9-sgRNA is a powerful tool for the exploration and manipulation of gene functions. Here we quantify the reversible switching of a central process of the bacterial cell cycle by CRISPRi and an antisense RNA mechanism. Reversible induction of filamentous growth in E. coli has been recently demonstrated by controlling the expression levels of the bacterial cell division proteins FtsZ/FtsA via CRISPRi. If FtsZ falls below a critical level, cells cannot divide. However, the cells remain metabolically active and continue with DNA replication. We surmised that this makes them amenable to an inducible antisense RNA strategy to counteract FtsZ inhibition. We show that both static and inducible thresholds can adjust the characteristics of the switching process. Combining bulk data with single cell measurements, we characterize the efficiency of the switching process. Successful restoration of division is found to occur faster in the presence of antisense sgRNAs than upon simple termination of CRISPRi induction.
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Extracellular vesicle-mediated delivery of CRISPR machinery silences androgen receptor in castration-resistant prostate cancer cells.
CRISPR-mediated gene editing is a promising technology for treatment of diseases by silencing a driver gene at the genomic DNA level. However, delivery of CRISPR machinery remains challenging for potential therapeutic application. Here, we developed a platform using extracellular vesicles (EVs) as a vehicle to deliver Cas9/single-guide RNA (sgRNA) ribonucleoprotein (RNP) complex to silence androgen receptor (AR) gene in prostate cancer (PCa) cells. A genetic modification conferred the N-myristoylation to the Cas9 protein, which enhanced the encapsulation of Cas9/sgRNA RNP into EVs and silenced both ectopic and endogenous AR gene. Interestingly, gene editing efficiency varied across PCa cell lines, associated with different chromatin accessibility at the target site. Functional analyses demonstrated that Cas9/sgRNA RNP (targeting the N-terminal domain of the AR gene) did not change gene-edited AR mRNA levels, but significantly inhibited expression levels of AR downstream genes, thereby attenuating PCa cell proliferation. Importantly, EV-mediated delivery of the Cas9/sgRNA RNP introduced indels into the AR gene and inhibited proliferation of enzalutamide-resistant PCa cells. This study highlights a therapeutic strategy for treatment of castration-resistant PCa using a programmable EV-mediated delivery platform.
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SgRNA Expression of CRIPSR-Cas9 System Based on MiRNA Polycistrons as a Versatile Tool to Manipulate Multiple and Tissue-Specific Genome Editing.
CRISPR/Cas9-mediated genome editing is a next-generation strategy for genetic modifications. Typically, sgRNA is constitutively expressed relying on RNA polymerase III promoters. Polymerase II promoters initiate transcription in a flexible manner, but sgRNAs generated by RNA polymerase II promoter lost their nuclease activity. To express sgRNAs in a tissue-specific fashion and endow CRISPR with more versatile function, a novel system was established in a polycistron, where miRNAs (or shRNAs) and sgRNAs alternately emerged and co-expressed under the control of a single polymerase II promoter. Effective expression and further processing of functional miRNAs and sgRNAs were achieved. The redundant nucleotides adjacent to sgRNA were degraded, and 5'- cap structure was responsible for the compromised nuclease capacity of sgRNA: Cas9 complex. Furthermore, this strategy fulfilled conducting multiplex genome editing, as well as executing neural- specific genome editing and enhancing the proportion of homologous recombination via inhibiting NHEJ pathway by shRNA. In summary, we designed a new construction for efficient expression of sgRNAs with miRNAs (shRNAs) by virtue of RNA polymerase II promoters, which will spur the development of safer, more controllable/regulable and powerful CRISPR/Cas9 system-mediated genome editing in a wide variety of further biomedical applications.
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Specific modulation of CRISPR transcriptional activators through RNA-sensing guide RNAs in mammalian cells and zebrafish embryos.
Cellular transcripts encode important information regarding cell identity and disease status. The activation of CRISPR in response to RNA biomarkers holds the potential for controlling CRISPR activity with spatiotemporal precision. This would enable the restriction of CRISPR activity to specific cell types expressing RNA biomarkers of interest while preventing unwanted activity in other cells. Here, we present a simple and specific platform for modulating CRISPR activity in response to RNA detection through engineering Streptococcus pyogenes Cas9 single-guide RNAs (sgRNAs). sgRNAs are engineered to fold into complex secondary structures that, in the ground state, inhibit their activity. Engineered sgRNAs become activated upon recognising complementary RNAs, thus enabling Cas9 to perform its function. Our approach enables CRISPR activation in response to RNA detection in both HEK293T cells and zebrafish embryos. Iterative design optimisations allowed the development of computational tools for generating sgRNAs capable of detecting RNA sequences of choice. Mechanistic investigations reveal that engineered sgRNAs are cleaved during RNA detection, and we identify key positions that benefit from chemical modifications to improve the stability of engineered sgRNAs in vivo. Our sensors open up novel opportunities for developing new research and therapeutic applications using CRISPR activation in response to endogenous RNA biomarkers.
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In vivo CRISPRa decreases seizures and rescues cognitive deficits in a rodent model of epilepsy.
Epilepsy is a major health burden, calling for new mechanistic insights and therapies. CRISPR-mediated gene editing shows promise to cure genetic pathologies, although hitherto it has mostly been applied ex vivo. Its translational potential for treating non-genetic pathologies is still unexplored. Furthermore, neurological diseases represent an important challenge for the application of CRISPR, because of the need in many cases to manipulate gene function of neurons in situ. A variant of CRISPR, CRISPRa, offers the possibility to modulate the expression of endogenous genes by directly targeting their promoters. We asked if this strategy can effectively treat acquired focal epilepsy, focusing on ion channels because their manipulation is known be effective in changing network hyperactivity and hypersynchronziation. We applied a doxycycline-inducible CRISPRa technology to increase the expression of the potassium channel gene Kcna1 (encoding Kv1.1) in mouse hippocampal excitatory neurons. CRISPRa-mediated Kv1.1 upregulation led to a substantial decrease in neuronal excitability. Continuous video-EEG telemetry showed that AAV9-mediated delivery of CRISPRa, upon doxycycline administration, decreased spontaneous generalized tonic-clonic seizures in a model of temporal lobe epilepsy, and rescued cognitive impairment and transcriptomic alterations associated with chronic epilepsy. The focal treatment minimizes concerns about off-target effects in other organs and brain areas. This study provides the proof-of-principle for a translational CRISPR-based approach to treat neurological diseases characterized by abnormal circuit excitability.
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Generation and validation of homozygous fluorescent knock-in cells using CRISPR-Cas9 genome editing.
Gene tagging with fluorescent proteins is essential for investigations of the dynamic properties of cellular proteins. CRISPR-Cas9 technology is a powerful tool for inserting fluorescent markers into all alleles of the gene of interest (GOI) and allows functionality and physiological expression of the fusion protein. It is essential to evaluate such genome-edited cell lines carefully in order to preclude off-target effects caused by (i) incorrect insertion of the fluorescent protein, (ii) perturbation of the fusion protein by the fluorescent proteins or (iii) nonspecific genomic DNA damage by CRISPR-Cas9. In this protocol, we provide a step-by-step description of our systematic pipeline to generate and validate homozygous fluorescent knock-in cell lines.We have used the paired Cas9D10A nickase approach to efficiently insert tags into specific genomic loci via homology-directed repair (HDR) with minimal off-target effects. It is time-consuming and costly to perform whole-genome sequencing of each cell clone to check for spontaneous genetic variations occurring in mammalian cell lines. Therefore, we have developed an efficient validation pipeline of the generated cell lines consisting of junction PCR, Southern blotting analysis, Sanger sequencing, microscopy, western blotting analysis and live-cell imaging for cell-cycle dynamics. This protocol takes between 6 and 9 weeks. With this protocol, up to 70% of the targeted genes can be tagged homozygously with fluorescent proteins, thus resulting in physiological levels and phenotypically functional expression of the fusion proteins.
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Accelerated genome engineering of Pseudomonas putida by I-SceI-mediated recombination and CRISPR-Cas9 counterselection.
Pseudomonas species have become reliable platforms for bioproduction due to their capability to tolerate harsh conditions imposed by large-scale bioprocesses and their remarkable resistance to diverse physicochemical stresses. The last few years have brought forth a variety of synthetic biology tools for the genetic manipulation of pseudomonads, but most of them are either applicable only to obtain certain types of mutations, lack efficiency, or are not easily accessible to be used in different Pseudomonas species (e.g. natural isolates). In this work, we describe a versatile, robust and user-friendly procedure that facilitates virtually any kind of genomic manipulation in Pseudomonas species in 3-5 days. The protocol presented here is based on DNA recombination forced by double-stranded DNA cuts (through the activity of the I-SceI homing meganuclease from yeast) followed by highly efficient counterselection of mutants (aided by a synthetic CRISPR-Cas9 device). The individual parts of the genome engineering toolbox, tailored for knocking genes in and out, have been standardized to enable portability and easy exchange of functional gene modules as needed. The applicability of the procedure is illustrated both by eliminating selected genomic regions in the platform strain P. putida KT2440 (including difficult-to-delete genes) and by integrating different reporter genes (comprising novel variants of fluorescent proteins) into a defined landing site in the target chromosome.
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