Name: development of knock-out muscle cell lines using lentivirus-mediated crispr/cas9 gene editing
Uploaded: 2022-06-16T14:30:00
Description: Watch this Scientific Journal Video about Development of Knock-Out Muscle Cell Lines using Lentivirus-Mediated CRISPR/Cas9 Gene Editing at JoVE.com

This work was funded by grants from Association Fran&#231;aise contre les myopathies (AFM-T&#233;l&#233;thon) and from Auvergne-Rh&#244;ne Alpes R&#233;gion (AURA).

Muscle biopsies were obtained from the Bank of Tissues for Research (Myobank, a partner in the EU network EuroBioBank, Paris, France) in accordance with European recommendations and French legislation. Written informed consent was obtained from all individuals. Immortalized myoblasts were kindly produced by Dr. V. Mouly (Myology Institute, Paris, France), and the protocols were approved by Myology Institute ethic committee (MESRI, n AC-2019-3502).
1. CRISPR guide design
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<ol>
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A major step on the way to the characterization of genes of unknown function involved in pathologies is the development of relevant cellular models to study the function of these genes. The use of gene editing using CRISPR/Cas9 is an exponentially growing field of research, and the development of knock-out models as presented here is among its most widely used applications. In this context, we propose here a versatile protocol to develop a human cell line knock-out in any gene of interest, allowing the characterization o...

With the progress of gene sequencing and the identification of mutations in genes of unknown functions in a specific tissue, the development of relevant cellular models to understand the function of a new target gene and confirm its involvement in the related pathophysiological mechanisms constitutes an essential tool. In addition, these models are of major importance for future therapeutic developments1,2, and constitute an interesting alternative to the development of knock-out animal models in straight line with the international recommendations for reduction in the use of animals in experimentation. Gene editing using CRISPR/Cas9 is among the most powerful tools currently available, which has allowed the development of many knock-out/knock-in models, and targeted gene validation using CRISPR/Cas9 is among the most widely used application of CRISPR/Cas93. The success of gene editing relies on the ability to introduce the CRISPR tools (the guide RNAs and the nuclease Cas9) in the target cell model, which can be a challenge in many difficult-to-transfect cells, such as muscle cells4. This challenge can be overcome with the use of virus, usually lentivirus, which has the great advantage to transduce efficiently many cell types and deliver its transgene. But its major drawback is the integration of the transgene in the host cell genome, leading to potential alteration of genes localized at the integration site and to the permanent expression of the transgene, which in the case of the nuclease Cas9 would result in damaging consequences5. A smart solution has been proposed by Merienne and colleagues6, which consists of introduction into the cells of a guide-RNA targeting the Cas9 gene itself, leading to Cas9 inactivation. An adaptation of this strategy is presented here as a user friendly and versatile protocol allowing to knock-out virtually any gene in difficult-to-transfect cells.
The goal of the protocol presented here is to induce the inactivation of a gene of interest in immortalized muscle cells. It can be used to knock-out any gene of interest, in different types of immortalized cells. The protocol described here contains steps to design the guide RNAs and their cloning into lentiviral plasmids, to produce the CRISPR tools in lentiviral vectors, to transduce the cells with the different lentiviruses, and to clone the cells to produce a homogenous edited cell line.
Using this protocol, immortalized human skeletal muscle cells have been developed with deletion of the type 1 ryanodine receptor (RyR1), an essential calcium channel involved in intracellular calcium release and muscle contraction7. The knock-out (KO) of the gene has been confirmed at the protein level using Western blot, and at the functional level using calcium imaging.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Anti-CACNA1S antibody</td>
			<td>Sigma-Aldrich</td>
			<td>HPA048892</td>
			<td>Primary antibody</td>
		</tr>
		<tr>
			<td>Blp I</td>
			<td>NE BioLabs</td>
			<td>R0585S</td>
			<td>Restriction enzyme</td>
		</tr>
		<tr>
			<td>CalPhos Mammalian Transfection Kit</td>
			<td>Takara</td>
			<td>631312&#160;</td>
			<td>Transfection kit</td>
		</tr>
		<tr>
			<td>Easy blot anti Mouse IgG</td>
			<td>GeneTex</td>
			<td>GTX221667-01</td>
			<td>HRP secondary antibody</td>
		</tr>
		<tr>
			<td>Easy blot anti Rabbit IgG</td>
			<td>GeneTex</td>
			<td>GTX221666</td>
			<td>HRP secondary antibody</td>
		</tr>
		<tr>
			<td>Fluo-4 direct</td>
			<td>Molecular Probes</td>
			<td>F10472</td>
			<td>Calcium imaging</td>
		</tr>
		<tr>
			<td>GAPDH(14C10) Rabbit mAb&#160;</td>
			<td>Cell Signaling Technology</td>
			<td>#2118</td>
			<td>Primary antibody</td>
		</tr>
		<tr>
			<td>HindIII</td>
			<td>Fermentas</td>
			<td>ER0501</td>
			<td>Restriction enzyme</td>
		</tr>
		<tr>
			<td>InFusion HD Precision Plus</td>
			<td>Takara</td>
			<td>638920</td>
			<td>Ligation kit</td>
		</tr>
		<tr>
			<td>MasterMix Phusion High Fidelity with GC</td>
			<td>ThermoFisher Scientific</td>
			<td>F532L</td>
			<td>Mix for PCR reaction with High fidelity Taq polymerase and dNTPs</td>
		</tr>
		<tr>
			<td>Myosin Heavy Chain antibody</td>
			<td>DHSB</td>
			<td>MF20</td>
			<td>Primary antibody</td>
		</tr>
		<tr>
			<td>NucleoBond Xtra Maxi EF</td>
			<td>Macherey-Nagel</td>
			<td>REF&#160;740424</td>
			<td>Maxipreparation kit for purification of plasmids</td>
		</tr>
		<tr>
			<td>NucleoSpin Gel and PCR Clean-up</td>
			<td>Macherey-Nagel</td>
			<td>740609</td>
			<td>DNA purification</td>
		</tr>
		<tr>
			<td>NucleoSpin Tissue</td>
			<td>Macherey-Nagel</td>
			<td>740952</td>
			<td>Kit for DNA extraction from cell</td>
		</tr>
		<tr>
			<td>One Shot Stbl3 Chemically Competent&#160;E. coli</td>
			<td>ThermoFisher Scientific</td>
			<td>C737303</td>
			<td>Chemically competent cells</td>
		</tr>
		<tr>
			<td>Plasmid #87904</td>
			<td>Addgene</td>
			<td>87904</td>
			<td>Lentiviral plasmid encoding the SpCas9 (for LV-Cas9)</td>
		</tr>
		<tr>
			<td>Plasmid #87919</td>
			<td>Addgene</td>
			<td>87919</td>
			<td>Lentiviral backbone for insertion of cassette with guides (for LV-guide-target)</td>
		</tr>
		<tr>
			<td>Plasmid #12260</td>
			<td>Addgene</td>
			<td>12260</td>
			<td>Lentiviral plasmid encoding lentiviral packaging GAG POL</td>
		</tr>
		<tr>
			<td>Plasmid #8454</td>
			<td>Addgene</td>
			<td>8454</td>
			<td>Lentiviral plasmid encoding envelope protein for producing lentiviral and MuLV retroviral particles</td>
		</tr>
		<tr>
			<td>V5 Tag Monoclonal Antibody</td>
			<td>Invitrogene</td>
			<td>R96025&#160;</td>
			<td>Primary antibody</td>
		</tr>
		<tr>
			<td>XL10-Gold Ultracompetent Cells</td>
			<td>Agilent</td>
			<td>200317</td>
			<td>Chemically competent cells</td>
		</tr>
		<tr>
			<td>Xma I</td>
			<td>NE BioLabs</td>
			<td>R0180S</td>
			<td>Restriction enzyme</td>
		</tr>
	</tbody>
</table>

development of knock-out muscle cell lines using lentivirus-mediated crispr/cas9 gene editing

One important application of clustered regulatory interspaced short palindromic repeats (CRISPR)/Cas 9 is the development of knock-out cell lines, specifically to study the function of new genes/proteins associated with a disease, identified during the genetic diagnosis. For the development of such cell lines, two major issues have to be untangled: insertion of the CRISPR tools (the Cas9 and the guide RNA) with high efficiency into the chosen cells, and restriction of the Cas9 activity to the specific deletion of the chosen gene. The protocol described here is dedicated to the insertion of the CRISPR tools in difficult to transfect cells, such as muscle cells. This protocol is based on the use of lentiviruses, produced with plasmids publicly available, for which all the cloning steps are described to target a gene of interest. The control of Cas9 activity has been performed using an adaptation of a previously described system called KamiCas9, in which the transduction of the cells with a lentivirus encoding a guide RNA targeting the Cas9 allows the progressive abolition of Cas9 expression. This protocol has been applied to the development of a RYR1-knock out human muscle cell line, which has been further characterized at the protein and functional level, to confirm the knockout of this important calcium channel involved in muscle intracellular calcium release and in excitation-contraction coupling. The procedure described here can easily be applied to other genes in muscle cells or in other difficult to transfect cells and produce valuable tools to study these genes in human cells.

This protocol was applied to immortalized myoblasts from a healthy subject15 (so-called HM cells, for human myoblasts), in which the RyR1 has been previously characterized16, in order to knock out the RYR1 gene encoding the RyR1 protein. The design of the guides RNA was made to delete the sequence encompassing part of exon 101 and intron 101 of the gene. Deletion of part of exon 101 is foreseen to result in disruption of the reading frame. In addition, exon 101 enc...

Watch this Scientific Journal Video about Development of Knock-Out Muscle Cell Lines using Lentivirus-Mediated CRISPR/Cas9 Gene Editing at JoVE.com

Development of Knock-Out Muscle Cell Lines using Lentivirus-Mediated CRISPR/Cas9 Gene Editing

The protocol describes how to generate knock-out myoblasts using CRISPR/Cas9, starting from the design of guide-RNAs to the cellular cloning and characterization of the knock-out clones.

One important application of clustered regulatory interspaced short palindromic repeats (CRISPR)/Cas 9 is the development of knock-out cell lines, ...

This protocol allows the development of muscle cell knock-out in any gene, in order to further study the function of this gene in muscle. The advantage is that it's cheap and faster than the development of knock-out animals to study any gene. This technique can be used, for example, to study the involvement of a newly identified gene in a myopathy.If the technique described here is applied to iPS cells it can lead to the development of knock-out cells for any gene in any type of cells. To begin clustered regularly interspaced short palindromic repeats or CRISPR-mediated gene deletion, first use genome browser tools like ensembl. org to identify the target gene along with the DNA sequences on both sides of it.To get the list of the gene exons, first click on the transcript of the protein coding sequence and then on Exons. Next, click on Download sequence and select only Genomic sequence to download the full consensus sequence of the whole gene. Scroll the list of the exons and introns of the gene to select the targeted ones.For designing guide RNA, first select nucleotide sequences of the introns immediately upstream and downstream of the target sequence. Then go to a website such as crispr.tefor. net and use the selected sequences to design two guide RNAs, each 20 nucleotides long without the Protospacer Adjacent Motif separated by a few hundred base pairs.Next determine the reverse complement sequence for each guide RNA and order the primers. To construct guide RNA cassettes for plasmid cloning, first run a PCR with one set of primers using the recipe and program described in the text, then separate the PCR products in 1%agarose gel using Tris-borate-EDTA buffer. Next excise the 300-base pair fragment and purify using a kit.Similarly, after running another PCR with the other primer set, purify a 400-base pair long DNA fragment into 20 microliters of elution buffer. To insert the guide RNA cassettes into a lentiviral plasmid backbone, first set up a double digestion reaction of the plasmid with the restriction enzymes Xma 1 and Blp 1 as described in the text. After incubating the reaction tube at 37 degrees Celsius for one hour, incubate it at 65 degrees Celsius for 20 minutes, then run the digestion product in a 1%agarose gel and purify the plasmid of about 10 kilobase pairs using an appropriate kit.Also quantify the purified product by measuring the optical density. To ligate the guide RNA cassette with the plasmid, prepare a reaction mix with the purified guide RNA, the linearized plasmid DNA, enzyme, and water to a final volume of 10 microliters, then incubate the reaction tube at 50 degrees Celsius for 15 minutes to generate the final plasmid, called pGuide. For lentivirus production, first seed a 145-centimeter plate with 1 million HEK 293 cells in DMEM with high glucose and pyruvate supplemented with 10%FBS and 1%penicillin or streptomycin.Then grow the cells at 37 degrees Celsius for three days in a 5%carbon dioxide incubator. On the fourth day check the cells to ensure 60 to 65%confluency. Next prepare the transfection solution consisting of the plasmid of interest, the plasmid encoding the virus envelope, the lentiviral packaging plasmid psPAX2, and calcium phosphate.Adjust the final volume to 1000 microliters with water. Then add the transfection solution dropwise to one milliliter of 2X HBS under agitation and incubate at room temperature for at least 10 minutes. After that, add the solution dropwise to the cells.For a homogenous distribution of the transfection solution, gently tilt the plate in all directions, then incubate the cells at 37 degrees Celsius in a 5%carbon dioxide incubator for at least five hours. Next, remove the media from the plates and wash the cells with PBS to get rid of the transfection reagents, then add 12 milliliters of fresh medium. After 48 hours of incubation at 37 degrees Celsius, pool the media from all the plates into a tube.Put the tube in a sealed bucket and centrifuge at 800 times G for five minutes at four degrees Celsius, then filter the supernatant using a 0.45-micrometer filter and centrifuge the filtrate at 68, 300 times G for two hours at four degrees Celsius, using a swinging bucket rotor. After removing the supernatant, keep the tubes upside down on a paper towel in a safety cabinet for five to 10 minutes for maximum removal of liquid from the pellets. Add 100 microliters of HEC proliferation medium and keep the tubes at four degrees Celsius for at least two hours.Then resuspend the pellet by pipetting. After collecting all the resuspended pellets, make aliquots of 10 or 25 microliters, then store the aliquots at minus 80 degrees Celsius after snap freezing in liquid hydrogen. For myoblast transduction, first seed each well of a 96-well plate with 100 microliters of proliferation medium containing 10, 000 myoblast cells.The next day, add precalculated volumes of LV guides and LV Cas9 to the cells in a safety cabinet. After five days of incubation, release the cells from the well by adding trypsin. After counting the cell numbers, transfer the cells from wells with 40 to 50%confluency to a new plate.After five hours of incubation, add calculated volumes of LV killer at a multiplicity of infection of 20 and incubate for five to 10 days or at least two passages in between. For functional characterization of the gene-edited clones by calcium imaging, plate 50, 000 cells in the center of a laminin-coated 35-millimeter dish. To induce differentiation, add the differentiation medium as described in the text.After six days of incubation, take off the culture medium and delineate the cells with a hydrophobic pen. Then add 50 microliters of Fluo-4 Direct, diluted one-to-one in differentiation medium, to the differentiated myotubes. After incubating the dishes at 37 degrees Celsius for 30 minutes, wash the cells twice with Krebs buffer supplemented with one milligram per milliliter of glucose.Next use an inverted fluorescence microscope or a confocal microscope with a 10X objective to measure the fluorescence variations at a rate of one frame per second for 90 seconds and choose a field with at least 10 myotubes. After removing the additional Krebs buffer, stimulate the cells with two milliliters of potassium chloride for membrane depolarization or two milliliters of florochloral metacresol or 4CmC or RyR1 direct stimulation. Visualize the fluorescence variation after stimulation, then quantify the fluorescence variation in each myotube.This protocol could be successfully used to knockout the gene RyR1 from human myoblasts, which resulted in shorter genomic DNA in the cells. RyR1 knock-out was also confirmed at the protein level as the RyR1 protein could not be detected in the targeted cells by Western blotting. The absence of RyR1 activity in myotubes differentiated from the RyR1 knock-out cells was further verified by Fluo-4 calcium imaging after direct RyR1 stimulation by 4CmC or indirect stimulation by potassium chloride-induced membrane depolarization.The sequence that will be deleted should be required to the prediction of a functional protein. This technique is ideal for the development of post-genomic tools to study the involvement of new gene in muscle disease.

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Genetics

QTL Mapping and CRISPR/Cas9 Editing to Identify a Drug Resistance Gene in Toxoplasma gondii

Scientific knowledge is intrinsically linked to available technologies and methods. This article will present two methods that allowed for the identification and verification of a drug resistance gene in the Apicomplexan parasite Toxoplasma gondii, the method of Quantitative Trait Locus (QTL) mapping using a Whole Genome Sequence (WGS) -based genetic map and the method of Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9 -based gene editing. The approach of QTL mapping allows one to test if there is a correlation between a genomic region(s) and a phenotype. Two datasets are required to run a QTL scan, a genetic map based on the progeny of a recombinant cross and a quantifiable phenotype assessed in each of the progeny of that cross. These datasets are then formatted to be compatible with R/qtl software that generates a QTL scan to identify significant loci correlated with the phenotype. Although this can greatly narrow the search window of possible candidates, QTLs span regions containing a number of genes from which the causal gene needs to be identified. Having WGS of the progeny was critical to identify the causal drug resistance mutation at the gene level. Once identified, the candidate mutation can be verified by genetic manipulation of drug sensitive parasites. The most facile and efficient method to genetically modify T. gondii is the CRISPR/Cas9 system. This system comprised of just 2 components both encoded on a single plasmid, a single guide RNA (gRNA) containing a 20 bp sequence complementary to the genomic target and the Cas9 endonuclease that generates a double-strand DNA break (DSB) at the target, repair of which allows for insertion or deletion of sequences around the break site. This article provides detailed protocols to use CRISPR/Cas9 based genome editing tools to verify the gene responsible for sinefungin resistance and to construct transgenic parasites.

QTL Mapping and CRISPR/Cas9 Editing to Identify a Drug Resistance Gene in Toxoplasma gondii

Details are presented on how QTL mapping with a whole genome sequence based genetic map can be used to identify a drug resistance gene in Toxoplasma gondii and how this can be verified with the CRISPR/Cas9 system that efficiently edits a genomic target, in this case the drug resistance gene.

Scientific knowledge is intrinsically linked to available technologies and methods. This article will present two methods that allowed for the ...

Selection-dependent and Independent Generation of CRISPR/Cas9-mediated Gene Knockouts in Mammalian Cells

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide RNA (sgRNA) that confers specificity, the Cas9 protein cleaves both DNA strands at the targeted locus. The DNA break can trigger either non-homologous end joining (NHEJ) or homology directed repair (HDR). NHEJ can introduce small deletions or insertions which lead to frame-shift mutations, while HDR allows for larger and more precise perturbations. Here, we present protocols for generating knockout cell lines by coupling established CRISPR/Cas9 methods with two options for downstream selection/screening. The NHEJ approach uses a single sgRNA cut site and selection-independent screening, where protein production is assessed by dot immunoblot in a high-throughput manner. The HDR approach uses two sgRNA cut sites that span the gene of interest. Together with a provided HDR template, this method can achieve deletion of tens of kb, aided by the inserted selectable resistance marker. The appropriate applications and advantages of each method are discussed.

Recent advances in the ability to genetically manipulate somatic cell lines hold great potential for basic and applied research. Here, we present two approaches for CRISPR/Cas9 generated knockout production and screening in mammalian cell lines, with and without the use of selectable markers.

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide ...

A Standard Methodology to Examine On-site Mutagenicity As a Function of Point Mutation Repair Catalyzed by CRISPR/Cas9 and SsODN in Human Cells

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point mutations, which often are responsible for a variety of human inherited disorders. Using a well-established cell-based model system, the point mutation of a single-copy mutant eGFP gene integrated into HCT116 cells has been repaired using this combinatorial approach. The analysis of corrected and uncorrected cells reveals both the precision of gene editing and the development of genetic lesions, when indels are created in uncorrected cells in the DNA sequence surrounding the target site. Here, the specific methodology used to analyze this combinatorial approach to the gene editing of a point mutation, coupled with a detailed experimental strategy to measuring indel formation at the target site, is outlined. This protocol outlines a foundational approach and workflow for investigations aimed at developing CRISPR/Cas9-based gene editing for human therapy. The conclusion of this work is that on-site mutagenesis takes place as a result of CRISPR/Cas9 activity during the process of point mutation repair. This work puts in place a standardized methodology to identify the degree of mutagenesis, which should be an important and critical aspect of any approach destined for clinical implementation.

This protocol outlines the workflow of a CRISPR/Cas9-based gene editing system for the repair of point mutations in mammalian cells. Here, we use a combinatorial approach to gene editing with a detailed follow-on experimental strategy for measuring indel formation at the target site&#8212;in essence, analyzing onsite mutagenesis.

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point ...

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

The complete genome of the channel catfish, Ictalurus punctatus, has been sequenced, leading to greater opportunities for studying channel catfish gene function. Gene knockout has been used to study these gene functions in vivo. The clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/Cas9) system is a powerful tool used to edit genomic DNA sequences to alter gene function. While the traditional approach has been to introduce CRISPR/Cas9 mRNA into the single cell embryos through microinjection, this can be a slow and inefficient process in catfish. Here, a detailed protocol for microinjection of channel catfish embryos with CRISPR/Cas9 protein is described. Briefly, eggs and sperm were collected and then artificial fertilization performed. Fertilized eggs were transferred to a Petri dish containing Holtfreter's solution. Injection volume was calibrated and then guide RNAs/Cas9 targeting the toll/interleukin 1 receptor domain-containing adapter molecule (TICAM 1) gene and rhamnose binding lectin (RBL) gene were microinjected into the yolk of one-cell embryos. The gene knockout was successful as indels were confirmed by DNA sequencing. The predicted protein sequence alterations due to these mutations included frameshift and truncated protein due to premature stop codons.

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

A simple and efficient microinjection protocol for gene editing in channel catfish embryos using the CRISPR/Cas9 system is presented. In this protocol, guide RNAs and Cas9 protein were microinjected into the yolk of one-cell embryos. This protocol has been validated by knocking out two channel catfish immune-related genes.

The complete genome of the channel catfish, Ictalurus punctatus, has been sequenced, leading to greater opportunities for studying channel catfish gene ...

Highly Efficient Gene Disruption of Murine and Human Hematopoietic Progenitor Cells by CRISPR/Cas9

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. Complete gene ablation in HSCs required the generation of knockout mice from which HSCs could be isolated, and gene ablation in primary human HSCs was not possible. Viral transduction could be used for knock-down approaches, but these suffered from variable efficacy. In general, genetic manipulation of human and mouse hematopoietic cells was hampered by low efficiencies and extensive time and cost commitments. Recently, CRISPR/Cas9 has dramatically expanded the ability to engineer the DNA of mammalian cells. However, the application of CRISPR/Cas9 to hematopoietic cells has been challenging, mainly due to their low transfection efficiencies, the toxicity of plasmid-based approaches and the slow turnaround time of virus-based protocols.
A rapid method to perform CRISPR/Cas9-mediated gene editing in murine and human hematopoietic stem and progenitor cells with knockout efficiencies of up to 90% is provided in this article. This approach utilizes a ribonucleoprotein (RNP) delivery strategy with a streamlined three-day workflow. The use of Cas9-sgRNA RNP allows for a hit-and-run approach, introducing no exogenous DNA sequences in the genome of edited cells and reducing off-target effects. The RNP-based method is fast and straightforward: it does not require cloning of sgRNAs, virus preparation or specific sgRNA chemical modification. With this protocol, scientists should be able to successfully generate knockouts of a gene of interest in primary hematopoietic cells within a week, including downtimes for oligonucleotide synthesis. This approach will allow a much broader group of users to adapt this protocol for their needs.

A protocol for fast CRISPR/Cas9-mediated gene disruption in mouse and human primary hematopoietic cells is described in this article. Cas9-sgRNA ribonucleoproteins are introduced via electroporation with sgRNAs generated through in vitro transcription and commercial Cas9. High editing efficiencies are achieved with limited time and financial cost.

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. ...

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

Malaria is a significant cause of morbidity and mortality worldwide. This disease, which primarily affects those living in tropical and subtropical regions, is caused by infection with Plasmodium parasites. The development of more effective drugs to combat malaria can be accelerated by improving our understanding of the biology of this complex parasite. Genetic manipulation of these parasites is key to understanding their biology; however, historically the genome of P. falciparum has been difficult to manipulate. Recently, CRISPR/Cas9 genome editing has been utilized in malaria parasites, allowing for easier protein tagging, generation of conditional protein knockdowns, and deletion of genes. CRISPR/Cas9 genome editing has proven to be a powerful tool for advancing the field of malaria research. Here, we describe a CRISPR/Cas9 method for generating glmS-based conditional knockdown mutants in P. falciparum. This method is highly adaptable to other types of genetic manipulations, including protein tagging and gene knockouts.

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

We describe a method for generating glmS-based conditional knockdown mutants in Plasmodium falciparum using CRISPR/Cas9 genome editing.

Malaria is a significant cause of morbidity and mortality worldwide. This disease, which primarily affects those living in tropical and subtropical ...

Endogenous Protein Tagging in Human Induced Pluripotent Stem Cells Using CRISPR/Cas9

A protocol is presented for generating human induced pluripotent stem cells (hiPSCs) that express endogenous proteins fused to in-frame&#160;N- or C-terminal fluorescent tags. The prokaryotic CRISPR/Cas9 system (clustered regularly interspaced short palindromic repeats/CRISPR-associated 9) may be used to introduce large exogenous sequences into genomic loci via homology directed repair (HDR). To achieve the desired knock-in, this protocol employs the ribonucleoprotein (RNP)-based approach where wild type Streptococcus pyogenes Cas9 protein, synthetic 2-part guide RNA (gRNA), and a donor template plasmid are delivered to the cells via electroporation. Putatively edited cells expressing the fluorescently tagged proteins are enriched by fluorescence activated cell sorting (FACS). Clonal lines are then generated and can be analyzed for precise editing outcomes. By introducing the fluorescent tag at the genomic locus of the gene of interest, the resulting subcellular localization and dynamics of the fusion protein can be studied under endogenous regulatory control, a key improvement over conventional overexpression systems. The use of hiPSCs as a model system for gene tagging provides the opportunity to study the tagged proteins in diploid, nontransformed cells. Since hiPSCs can be differentiated into multiple cell types, this approach provides the opportunity to create and study tagged proteins in a variety of isogenic cellular contexts.

Described here is a protocol for tagging endogenously expressed proteins with fluorescent tags in human induced pluripotent stem cells using CRISPR/Cas9. Putatively edited cells are enriched by fluorescence activated cell sorting and clonal cell lines are generated.

A protocol is presented for generating human induced pluripotent stem cells (hiPSCs) that express endogenous proteins fused to in-frame&#160;N- or ...

Genome Editing in Mammalian Cell Lines using CRISPR-Cas

The clustered regularly interspaced short palindromic repeats (CRISPR) system functions naturally in bacterial adaptive immunity, but has been successfully repurposed for genome engineering in many different living organisms. Most commonly, the wildtype CRISPR associated 9 (Cas9) or Cas12a endonuclease is used to cleave specific sites in the genome, after which the DNA double-stranded break is repaired via the non-homologous end joining (NHEJ) pathway or the homology-directed repair (HDR) pathway depending on whether a donor template is absent or present respectively. To date, CRISPR systems from different bacterial species have been shown to be capable of performing genome editing in mammalian cells. However, despite the apparent simplicity of the technology, multiple design parameters need to be considered, which often leave users perplexed about how best to carry out their genome editing experiments. Here, we describe a complete workflow from experimental design to identification of cell clones that carry desired DNA modifications, with the goal of facilitating successful execution of genome editing experiments in mammalian cell lines. We highlight key considerations for users to take note of, including the choice of CRISPR system, the spacer length, and the design of a single-stranded oligodeoxynucleotide (ssODN) donor template. We envision that this workflow will be useful for gene knockout studies, disease modeling efforts, or the generation of reporter cell lines.

CRISPR-Cas is a powerful technology to engineer the complex genomes of plants and animals. Here, we detail a protocol to efficiently edit the human genome using different Cas endonucleases. We highlight important considerations and design parameters to optimize editing efficiency.

The clustered regularly interspaced short palindromic repeats (CRISPR) system functions naturally in bacterial adaptive immunity, but has been ...

CRISPR/Cas9 Ribonucleoprotein-mediated Precise Gene Editing by Tube Electroporation

Gene editing nucleases, represented by CRISPR-associated protein 9 (Cas9), are becoming mainstream tools in biomedical research. Successful delivery of CRISPR/Cas9 elements into the target cells by transfection is a prerequisite for efficient gene editing. This protocol demonstrates that tube electroporation (TE) machine-mediated delivery of CRISPR/Cas9 ribonucleoprotein (RNP), along with single-stranded oligodeoxynucleotide (ssODN) donor templates to different types of mammalian cells, leads to robust precise gene editing events. First, TE was applied to deliver CRISPR/Cas9 RNP and ssODNs to induce disease-causing mutations in the interleukin 2 receptor subunit gamma (IL2RG) gene and sepiapterin reductase (SPR) gene in rabbit fibroblast cells. Precise mutation rates of 3.57%-20% were achieved as determined by bacterial TA cloning sequencing. The same strategy was then used in human iPSCs on several clinically relevant genes including epidermal growth factor receptor (EGFR), myosin binding protein C, cardiac (Mybpc3), and hemoglobin subunit beta (HBB). Consistently, highly precise mutation rates were achieved (11.65%-37.92%) as determined by deep sequencing (DeepSeq). The present work demonstrates that tube electroporation of CRISPR/Cas9 RNP represents an efficient transfection protocol for gene editing in mammalian cells.

Presented here is a protocol for efficient CRISPR/Cas9 ribonucleoprotein-mediated gene editing in mammalian cells using tube electroporation.

Gene editing nucleases, represented by CRISPR-associated protein 9 (Cas9), are becoming mainstream tools in biomedical research. Successful delivery of ...

Direct Gene Knock-out of Axolotl Spinal Cord Neural Stem Cells via Electroporation of CAS9 Protein-gRNA Complexes

The axolotl has the unique ability to fully regenerate its spinal cord. This is largely due to the ependymal cells remaining as neural stem cells (NSCs) throughout life, which proliferate to reform the ependymal tube and differentiate into lost neurons after spinal cord injury. Deciphering how these NSCs retain pluripotency post-development and proliferate upon spinal cord injury to reform the exact pre-injury structure can provide valuable insight into how mammalian spinal cords may regenerate as well as potential treatment options. Performing gene knock-outs in specific subsets of NSCs within a restricted time period will allow study of the molecular mechanisms behind these regenerative processes, without being confounded by development perturbing effects. Described here is a method to perform gene knock-out in axolotl spinal cord NSCs using the CRISPR-Cas9 system. By injecting the CAS9-gRNA complex into the spinal cord central canal followed by electroporation, target genes are knocked out in NSCs within specific regions of the spinal cord at a desired timepoint, allowing for molecular studies of spinal cord NSCs during regeneration.

Presented here is a protocol to perform time- and space-restricted gene knock-out in axolotl spinal cords by injecting CAS9-gRNA complex into the spinal cord central canal followed by electroporation.

The axolotl has the unique ability to fully regenerate its spinal cord. This is largely due to the ependymal cells remaining as neural stem cells (NSCs) ...