This work was supported by the Wellcome Trust grant number 206194 awarded to GJW. We thank the Cytometry Core facility: Bee Ling Ng, Jennifer Graham, Sam Thompson, and Christopher Hall for help with FACS.

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The authors have nothing to disclose.

A CRISPR-based screening strategy to identify genes encoding cellular components involved in cellular recognition is described. A similar approach using CRISPR activation also provides a genetic alternative to identify directly interacting receptors of recombinant proteins without the need to generate large protein libraries26. However, one major advantage of this approach is that it is applicable to interactions mediated by surface molecules natively displayed on the cell and does not depend on the overexpression of receptors, which can influence the binding avidity of the receptor. Unlike other methods, therefore, this technique makes no assumptions regarding the biochemical nature or cell biology of the receptors and provides an opportunity to study interactions mediated by proteins that are normally difficult to study using biochemical approaches, such as very large proteins, or those that traverse the membrane multiple times or form complexes with other proteins, and molecules other than proteins such as glycans, glycolipids, and phospholipids. Given the genome-scale nature of the method, this approach also has the advantage of not only identifying the receptor but also additional cellular components that are required for the binding event, thereby providing insights into the cell biology of the receptor.
One of the major limitations of this method when using it to identify the receptor of an orphan protein is the initial requirement to first identify a cell line that binds to the protein. This is not always easy and identifying a cell line that displays a binding phenotype that is also permissive to genetic screens can be the time-limiting step for deploying this assay. Some cell lines tend to bind to more proteins than others. This is especially relevant for proteins that bind HS, because these proteins tend to bind to any cell line that displays HS side chains, irrespective of the native binding context. Additionally, we have observed that upregulation of syndecans (i.e., proteoglycans that contain HS) in cell lines leads to increased binding of HS-binding proteins26. This could be a factor to take into consideration when selecting the cell line for screening. However, also important to note is that the additive binding of HS does not interfere with the binding to a specific receptor. This means that if binding is observed, it is possible that it is mediated solely by HS because the binding mediated by HS in this assay is additive rather than codependent19. In such a scenario, the heparin blocking approach described can identify such behaviors without having the need to perform a full genetic screen.
A useful resource for choosing cell lines is Cell Model Passport, which contains genomics, transcriptomics, and culture condition information for ~1,000 cancer cell lines27. Depending on the biological context, cells can be chosen based on their expression profiles. To aid the selection of cell lines, we clustered ~1,000 cell lines in Cell Model Passport according to the expression of ~1,500 preannotated human cell surface glycoproteins28 (Supplementary Figure 2; cluster information for each cell line together with growth conditions are provided in Supplementary Table 2). When testing the binding of a protein with unknown function, it is useful to select a panel of representative cell lines from each cluster to increase the chance of covering a wide range of receptors. Given a choice, it is recommended to choose cell lines that are easy to culture and easy to transduce. As these cell lines will be used in genome-scale screening, it is preferable that they can be grown easily in large quantities and are permissive to lentiviral transduction, because it is the most commonly available method for delivery of sgRNA for CRISPR-based genetic screening in the later steps.
Generally, the phenotype selections are carried out in a single sort. However, this is determined by the brightness of the stained cell population compared to the control. Iterative rounds of selections could be adopted for scenarios in which the signal-to-noise ratio of the desired phenotype is low, or when the aim of the screen is to identify mutants that have strong phenotypes. When using an iterative selection approach for FACS-based screens, it is important to consider that the sorting process can cause cell death, mainly due to the sheer force of the sorter. Thus, not all collected cells will be represented in the next round of sorting.
Library complexity is a very important factor in performing successful genetic screens, especially for negative selection screens because the extent of depletion in these can only be determined by comparing results to what was present in the starting library. For negative selection screens, it is common to maintain libraries of 500-1,000 x complexity. Positive selection screens, however, are more robust to library sizes, because in such screens only a small number of mutants are expected to be selected for a particular phenotype. Therefore, in the positive selection screen described here, the library size can be decreased to 50-100x complexity without compromising the quality of the screen. In addition, in these screens it is also possible to use a control library for a given cell line on a given day as a &#34;general control&#34; for all samples sorted on the day for that given cell line. This will reduce the number of control libraries that need to be produced and sequenced.
Another important consideration for using this approach is the limitations of loss-of-function screens in identifying genes that are essential for in vitro cell growth. The timing of the screens is crucial in this regard, as the longer the mutant cells are kept in culture, the higher the likelihood that cells with mutations in essential genes become nonviable and are no longer represented in the mutant library. The recent genetic screens performed as a part of the Project Score initiative in over 300 cell lines show that multiple genes in the KEGG-annotated protein secretion and N-glycosylation pathway are often identified as being essential for a number of cell lines (Supplementary Figure 3)29. This can be taken into consideration when designing screens if the effect of genes required for proliferation and viability is to be investigated in the context of cellular recognition process. In this case, carrying out screens at an early timepoint (e.g., day 9 posttransduction) would be generally appropriate. However, if the approach is used to identify a few targets with strong size effects rather than general cellular pathways, it might be appropriate to perform screens at a later time point (e.g., day 15-16 posttransduction).
The results from the screening are very robust; in eight recombinant protein binding screens performed in the past, the cell surface receptor was the top hit in every case19. When using this approach to identify the interaction partner, one should therefore expect the receptor and the factors contributing to its presentation on the surface to be identified with a high statistical confidence. Once the screen is performed and a hit is validated using a single gRNA knockout, further follow-ups can be performed using existing biochemical methods such as AVEXIS4 and direct saturable binding of purified proteins using surface plasmon resonance. The approach described here is applicable for all proteins for which it is possible to generate a soluble recombinant binding probe.
In summary, this is a genome-scale CRISPR knockout approach to identify interactions mediated by cell surface proteins. This method is generally applicable to identify cellular pathways required for cell surface recognition in a wide range of different biological contexts, including between an organism&#39;s own cells (e.g., neural and immunological recognition), as well as between host cells and pathogen proteins. This method provides a genetic alternative to biochemical approaches designed for receptor identification, and because it does not require any prior assumptions regarding the biochemical nature or cell biology of the receptors it has great potential to make completely unexpected discoveries.

Extracellular interactions by cell surface receptor proteins direct important biological processes such as tissue organization, host-pathogen recognition, and immune regulation. Investigating these interactions is of interest to the wider biomedical community, because membrane receptors are actionable targets of systematically delivered therapeutics such as monoclonal antibodies. Despite their importance, studying these interactions remains technically challenging. This is mainly because membrane-embedded receptors are amphipathic, making them difficult to isolate from biological membranes for biochemical manipulation, and their interactions are typified by the weak interaction affinities (KDs in the &#181;M-mM range)1. Consequently, many commonly used methods are unsuitable to detect this class of protein interactions1,2.
A range of methods has been developed to specifically investigate extracellular receptor-ligand interactions that take their unique biochemical properties into consideration3. A number of these approaches involve expressing the entire ectodomain of a receptor as a soluble recombinant protein in mammalian or insect cell-based systems to ensure that these proteins contain posttranslational modifications that are structurally important, such as glycans and disulfide bonds. To overcome the low-affinity binding, the ectodomains are often oligomerized to increase their binding avidity. Avid protein ectodomains have been successfully used as binding probes to identify interaction partners in direct recombinant protein-protein interaction screens4,5,6,7. While broadly successful, recombinant protein-based methods require that the ectodomain of a membrane receptor be produced as a soluble protein. Therefore, it is only generally applicable to proteins that contain a contiguous extracellular region (e.g., single-pass type I, type II, or GPI-anchored) and is not generally suitable for receptor complexes and membrane proteins that span the membrane multiple times.
Expression cloning techniques in which a library of complementary DNAs (cDNAs) is transfected into cells and tested for a gain-of-binding phenotype have also been used to identify extracellular protein-protein interactions8. The availability of large collections of cloned and sequenced cDNA expression plasmids in recent years has facilitated methods in which cell lines overexpressing cDNAs encoding cell surface receptors are screened for the binding of recombinant proteins to identify interactions9,10. The cDNA overexpression-based approaches, unlike recombinant protein-based methods, afford the possibility to identify interactions in the context of the plasma membrane. However, the success of using cDNA expression constructs depends on the cells&#39; ability to overexpress the protein in the correctly folded form, but this often requires cellular accessory factors such as transporters, chaperones, and correct oligomeric assembly. Transfecting a single cDNA might therefore not be enough to achieve cell surface expression.
Screening techniques using cDNA constructs or recombinant protein probes are resource-intensive and require large collections of cDNA or recombinant protein libraries. Specifically designed mass spectrometry-based methods have been utilized recently to identify extracellular interactions that do not require the assembly of large libraries. However, these techniques require chemical manipulation of the cell surface, which can alter the biochemical nature of the molecules present on the surface of the cells and are currently only applicable for interactions mediated by glycosylated proteins11,12. The majority of the currently available methods also heavily focus on the interactions between proteins while largely ignoring the contribution from the membrane microenvironment, including molecules such as glycans, lipids, and cholesterol.
The recent development of highly efficient bialleleic targeting using CRISPR-based approaches has enabled genome-scale libraries of cells lacking defined genes in a single pool that can be screened in a systematic and unbiased way to identify cellular components involved in different contexts, including dissecting cellular signaling processes, identification of perturbations that confer resistance to drugs, toxins, and pathogens, and determining specificity of antibodies13,14,15,16. Here, we describe a genome-scale CRISPR-based knockout cell screening assay that provides an alternative to the current biochemical approaches to identify extracellular receptor-ligand interactions. This approach of identifying interactions mediated by membrane receptors by genetic screens is particularly suitable for researchers that have a focused interest on individual ligands because it avoids the need to compile large libraries of cDNAs or recombinant proteins.
This assay consists of three major steps: 1) Highly avid recombinant protein binding probes consisting of the extracellular regions of a receptor of interest are produced and used in fluorescence-based flow cytometry-based binding assays; 2) The binding assays are used to identify a cell line that expresses the interaction partner of the recombinant protein probe; 3) A Cas9-expressing version of the cell line that interacts with the protein of interest is produced and a genome-scale CRISPR/Cas9-based knockout screen is performed (Figure 1). In this genetic screen, binding of a recombinant protein to cell lines is used as a measurable phenotype in which cells within the knockout library that have lost the ability to bind the probe are sorted using fluorescence-based activated cell sorting (FACS) and the genes that caused the loss of the binding phenotype identified by sequencing. In principle, the genes encoding the receptor responsible for binding the avid probe and those required for its cell surface display are identified.
The first step of this protocol involves the production of avid recombinant protein probes representing the ectodomain of the membrane-bound receptors. These receptors are known to frequently retain their extracellular binding functions when their ectodomains are expressed as a recombinant soluble protein1. For a protein of interest, soluble recombinant proteins can be produced in any suitable eukaryotic protein expression system in any format provided that it can be oligomerized for increased binding avidity, and it contains tags that can be used in fluorescence-based flow cytometry-based binding assays (e.g., FLAG-tag, biotin-tag). Detailed protocols for the production of soluble ectodomains of membrane receptors using the HEK293 protein expression system, as well as different multimerization techniques and the protein expression constructs for the production of both pentameric proteins and monomeric proteins have been previously described1,17. The protocol here will describe the steps for generating fluorescent avid probes from monomeric biotinylated proteins by conjugating them to streptavidin conjugated to a fluorochrome (e.g., phycoerythrin, or PE), which can be used directly in cell-based binding assays and has the advantage of not requiring a secondary antibody for detection. General protocols for performing genome-scale screens have already been described20,21, thus the protocol mainly focus on the specifics of performing flow cytometry-based recombinant protein binding screens using the CRISPR/Cas9 knockout screening system using the Human V1 (&#34;Yusa&#34;) library18.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Anti-mouse alkaline phosphatase</td>
			<td>Sigma</td>
			<td>A4656</td>
			<td></td>
		</tr>
		<tr>
			<td>Blasticidin</td>
			<td>Chem-Cruz</td>
			<td>SC-204655</td>
			<td></td>
		</tr>
		<tr>
			<td>Blood &#38; Cell Culture DNA Maxi Kit</td>
			<td>Qiagen</td>
			<td>13362</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma</td>
			<td>A9647-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>Diethanolamine</td>
			<td>Sigma</td>
			<td>398179</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Gibco</td>
			<td>31966-021</td>
			<td></td>
		</tr>
		<tr>
			<td>Dneasy Blood and Tissue kit</td>
			<td>Qiagen</td>
			<td>69504</td>
			<td></td>
		</tr>
		<tr>
			<td>DynaMag-96 Side Magnet</td>
			<td>Invitrogen</td>
			<td>12331D</td>
			<td></td>
		</tr>
		<tr>
			<td>HEK293T packaging cells</td>
			<td>ATCC</td>
			<td>CRL-3216</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>Sigma</td>
			<td>H4784-1G</td>
			<td></td>
		</tr>
		<tr>
			<td>KAPA HiFi HotStart ReadyMix</td>
			<td>Kapa</td>
			<td>KK2602</td>
			<td></td>
		</tr>
		<tr>
			<td>Lipofectamine LTX with PLUS reagent</td>
			<td>Invitrogen</td>
			<td>15338100</td>
			<td></td>
		</tr>
		<tr>
			<td>MoFlo XDP cell sorter</td>
			<td>BD</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ni2+-NTA agarose beads</td>
			<td>Jena Bioscience</td>
			<td>AC-501-25</td>
			<td></td>
		</tr>
		<tr>
			<td>OPTI-MEM</td>
			<td>Life Technologies</td>
			<td>31985-070</td>
			<td></td>
		</tr>
		<tr>
			<td>OX-68 antibody</td>
			<td>AbD Serotec</td>
			<td>MCA1022R</td>
			<td></td>
		</tr>
		<tr>
			<td>p-nitrophenyl phosphate</td>
			<td>Sigma</td>
			<td>1040-506</td>
			<td></td>
		</tr>
		<tr>
			<td>PD-10 desalting columns</td>
			<td>GE healthcare</td>
			<td>17085101</td>
			<td></td>
		</tr>
		<tr>
			<td>Polybrene</td>
			<td>Millipore</td>
			<td>TR-1003-G</td>
			<td></td>
		</tr>
		<tr>
			<td>Polypropylene tubes with 5 mL bed volume</td>
			<td>Qiagen</td>
			<td>34964</td>
			<td></td>
		</tr>
		<tr>
			<td>Proteinase K, recombinant, PCR Grade</td>
			<td>Roche</td>
			<td>3115879001</td>
			<td></td>
		</tr>
		<tr>
			<td>Puromycin</td>
			<td>Gibco</td>
			<td>A11138-03</td>
			<td></td>
		</tr>
		<tr>
			<td>Q5 Hot Start High-Fidelity 2&#215; Master Mix</td>
			<td>NEB</td>
			<td>M0494L</td>
			<td></td>
		</tr>
		<tr>
			<td>QIAquick PCR purification kit</td>
			<td>Qiagen</td>
			<td>28104</td>
			<td></td>
		</tr>
		<tr>
			<td>SCFA filter</td>
			<td>Nalgene</td>
			<td>190-2545</td>
			<td></td>
		</tr>
		<tr>
			<td>Sony Cell sorter</td>
			<td>Sony</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>SPRI beads (Agencourt AMPure XP beads)</td>
			<td>Beckman</td>
			<td>A63881</td>
			<td></td>
		</tr>
		<tr>
			<td>Streptavidin-coated microtitre plates</td>
			<td>Nalgene</td>
			<td>734-1284</td>
			<td></td>
		</tr>
		<tr>
			<td>Streptavidin-PE</td>
			<td>Biolegend</td>
			<td>405204</td>
			<td></td>
		</tr>
	</tbody>
</table>

cell surface receptor identification using genome-scale crispr/cas9 genetic screens

Intercellular communication mediated by direct interactions between membrane-embedded cell surface receptors is crucial for the normal development and functioning of multicellular organisms. Detecting these interactions remains technically challenging, however. This manuscript describes a systematic genome-scale CRISPR/Cas9 knockout genetic screening approach that reveals cellular pathways required for specific cell surface recognition events. This assay utilizes recombinant proteins produced in a mammalian protein expression system as avid binding probes to identify interaction partners in a cell-based genetic screen. This method can be used to identify the genes necessary for cell surface interactions detected by recombinant binding probes corresponding to the ectodomains of membrane-embedded receptors. Importantly, given the genome-scale nature of this approach, it also has the advantage of not only identifying the direct receptor but also the cellular components that are required for the presentation of the receptor at the cell surface, thereby providing valuable insights into the biology of the receptor.

Watch this Scientific Journal Video about Cell Surface Receptor Identification Using Genome-Scale CRISPR/Cas9 Genetic Screens at JoVE.com

Cell Surface Receptor Identification Using Genome-Scale CRISPR/Cas9 Genetic Screens

This manuscript describes a genome-scale cell-based screening approach to identify extracellular receptor-ligand interactions.

Intercellular communication mediated by direct interactions between membrane-embedded cell surface receptors is crucial for the normal development and ...

cell-surface-receptor-identification-using-genome-scale-crisprcas9

Research

JoVE Journal

Genetics

14.3K Views.  Wellcome Trust Sanger Institute. This manuscript describes a genome-scale cell-based screening approach to identify extracellular receptor-ligand interactions.

Video: Cell Surface Receptor Identification Using Genome-Scale CRISPR/Cas9 Genetic Screens

Using a Fluorescent PCR-capillary Gel Electrophoresis Technique to Genotype CRISPR/Cas9-mediated Knockout Mutants in a High-throughput Format

The development of programmable genome-editing tools has facilitated the use of reverse genetics to understand the roles specific genomic sequences play in the functioning of cells and whole organisms. This cause has been tremendously aided by the recent introduction of the CRISPR/Cas9 system-a versatile tool that allows researchers to manipulate the genome and transcriptome in order to, among other things, knock out, knock down, or knock in genes in a targeted manner. For the purpose of knocking out a gene, CRISPR/Cas9-mediated double-strand breaks recruit the non-homologous end-joining DNA repair pathway to introduce the frameshift-causing insertion or deletion of nucleotides at the break site. However, an individual guide RNA may cause undesirable off-target effects, and to rule these out, the use of multiple guide RNAs is necessary. This multiplicity of targets also means that a high-volume screening of clones is required, which in turn begs the use of an efficient high-throughput technique to genotype the knockout clones. Current genotyping techniques either suffer from inherent limitations or incur high cost, hence rendering them unsuitable for high-throughput purposes. Here, we detail the protocol for using fluorescent PCR, which uses genomic DNA from crude cell lysate as a template, and then resolving the PCR fragments via capillary gel electrophoresis. This technique is accurate enough to differentiate one base-pair difference between fragments and hence is adequate in indicating the presence or absence of a frameshift in the coding sequence of the targeted gene. This precise knowledge effectively precludes the need for a confirmatory sequencing step and allows users to save time and cost in the process. Moreover, this technique has proven to be versatile in genotyping various mammalian cells of various tissue origins targeted by guide RNAs against numerous genes, as shown here and elsewhere.

The genotyping technique described here, which couples fluorescent polymerase chain reaction (PCR) to capillary gel electrophoresis, allows for high-throughput genotyping of nuclease-mediated knockout clones. It circumvents limitations faced by other genotyping techniques and is more cost effective than sequencing methods.

The development of programmable genome-editing tools has facilitated the use of reverse genetics to understand the roles specific genomic sequences play ...

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 ...

Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus

The creation of mutant lines by genome editing is accelerating genetic analysis in many organisms. CRISPR/Cas9 methods have been adapted for use in the African clawed frog, Xenopus, a longstanding model organism for biomedical research. Traditional breeding schemes for creating homozygous mutant lines with CRISPR/Cas9-targeted mutagenesis have several time-consuming and laborious steps. To facilitate the creation of mutant embryos, particularly to overcome the obstacles associated with knocking out genes that are essential for embryogenesis, a new method called leapfrogging was developed. This technique leverages the robustness of Xenopus embryos to &#34;cut and paste&#34; embryological methods. Leapfrogging utilizes the transfer of primordial germ cells (PGCs) from efficiently-mutagenized donor embryos into PGC-ablated wildtype siblings. This method allows for the efficient mutation of essential genes by creating chimeric animals with wildtype somatic cells that carry a mutant germline. When two F0 animals carrying &#34;leapfrog transplants&#34; (i.e., mutant germ cells) are intercrossed, they produce homozygous, or compound heterozygous, null F1 embryos, thus saving a full generation time to obtain phenotypic data. Leapfrogging also provides a new approach for analyzing maternal effect genes, which are refractory to F0 phenotypic analysis following CRISPR/Cas9 mutagenesis. This manuscript details the method of leapfrogging, with special emphasis on how to successfully perform PGC transplantation.

Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus

Genes essential for survival pose technical hurdles for creating mutant lines. Leapfrogging circumvents lethality by combining genome editing with primordial germ cell transplantation to create wild-type animals carrying germline mutations. Leapfrogging also permits the efficient generation of homozygous null mutants in the F1 generation. Here, the transplantation step is demonstrated.

The creation of mutant lines by genome editing is accelerating genetic analysis in many organisms. CRISPR/Cas9 methods have been adapted for use in the ...

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration of transgenes. However, due to the low efficiency of homologous recombination (HR) and various indel mutations of non-homologous end joining (NHEJ)-based strategies in non-dividing cells, in vivo genome editing remains a great challenge. Here, we describe a homology-mediated end joining (HMEJ)-based CRISPR/Cas9 system for efficient in vivo precise targeted integration. In this system, the targeted genome and the donor vector containing homology arms (~800 bp) flanked by single guide RNA (sgRNA) target sequences are cleaved by CRISPR/Cas9. This HMEJ-based strategy achieves efficient transgene integration in mouse zygotes, as well as in hepatocytes in vivo. Moreover, a HMEJ-based strategy offers an efficient approach for correction of fumarylacetoacetate hydrolase (Fah) mutation in the hepatocytes and rescues Fah-deficiency induced liver failure mice. Taken together, focusing on targeted integration, this HMEJ-based strategy provides a promising tool for a variety of applications, including generation of genetically modified animal models and targeted gene therapies.

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

The clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/Cas9) system provides a promising tool for genetic engineering, and opens up the possibility of targeted integration of transgenes. We describe a homology-mediated end joining (HMEJ)-based strategy for efficient DNA targeted integration in vivo and targeted gene therapies using CRISPR/Cas9.

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration ...

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development of a customizable platform to rapidly generate gene modifications in a wide variety of organisms, including zebrafish. CRISPR-based genome editing uses a single guide RNA (sgRNA) to target a CRISPR-associated (Cas) endonuclease to a genomic DNA (gDNA) target of interest, where the Cas endonuclease generates a double-strand break (DSB). Repair of DSBs by error-prone mechanisms lead to insertions and/or deletions (indels). This can cause frameshift mutations that often introduce a premature stop codon within the coding sequence, thus creating a protein-null allele. CRISPR-based genome engineering requires only a few molecular components and is easily introduced into zebrafish embryos by microinjection. This protocol describes the methods used to generate CRISPR reagents for zebrafish microinjection and to identify fish exhibiting germline transmission of CRISPR-modified genes. These methods include in vitro transcription of sgRNAs, microinjection of CRISPR reagents, identification of indels induced at the target site using a PCR-based method called a heteroduplex mobility assay (HMA), and characterization of the indels using both a low throughput and a powerful next-generation sequencing (NGS)-based approach that can analyze multiple PCR products collected from heterozygous fish. This protocol is streamlined to minimize both the number of fish required and the types of equipment needed to perform the analyses. Furthermore, this protocol is designed to be amenable for use by laboratory personal of all levels of experience including undergraduates, enabling this powerful tool to be economically employed by any research group interested in performing CRISPR-based genomic modification in zebrafish.

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Targeted genome editing in the model system Danio rerio (zebrafish) has been greatly facilitated by the emergence of CRISPR-based approaches. Herein, we describe a streamlined, robust protocol for generation and identification of CRISPR-derived nonsense alleles that incorporates the heteroduplex mobility assay and identification of mutations using next-generation sequencing.

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development ...

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. ...

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 ...

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 ...

Pooled CRISPR-Based Genetic Screens in Mammalian Cells

Genome editing using the CRISPR-Cas system has vastly advanced the ability to precisely edit the genomes of various organisms. In the context of mammalian cells, this technology represents a novel means to perform genome-wide genetic screens for functional genomics studies. Libraries of guide RNAs (sgRNA) targeting all open reading frames permit the facile generation of thousands of genetic perturbations in a single pool of cells that can be screened for specific phenotypes to implicate gene function and cellular processes in an unbiased and systematic way. CRISPR-Cas screens provide researchers with a simple, efficient, and inexpensive method to uncover the genetic blueprints for cellular phenotypes. Furthermore, differential analysis of screens performed in various cell lines and from different cancer types can identify genes that are contextually essential in tumor cells, revealing potential targets for specific anticancer therapies. Performing genome-wide screens in human cells can be daunting, as this involves the handling of tens of millions of cells and requires analysis of large sets of data. The details of these screens, such as cell line characterization, CRISPR library considerations, and understanding the limitations and capabilities of CRISPR technology during analysis, are often overlooked. Provided here is a detailed protocol for the successful performance of pooled genome-wide CRISPR-Cas9 based screens.

CRISPR-Cas9 technology provides an efficient method to precisely edit the mammalian genome in any cell type and represents a novel means to perform genome-wide genetic screens. A detailed protocol discussing the steps required for the successful performance of pooled genome-wide CRISPR-Cas9 screens is provided here.

Genome editing using the CRISPR-Cas system has vastly advanced the ability to precisely edit the genomes of various organisms. In the context of mammalian ...

Introducing a Gene Knockout Directly Into the Amastigote Stage of Trypanosoma cruzi Using the CRISPR/Cas9 System

Trypanosoma cruzi is a pathogenic protozoan parasite that causes Chagas&#8217; disease mainly in Latin America. In order to identify a novel drug target against T. cruzi, it is important to validate the essentiality of the target gene in the mammalian stage of the parasite, the amastigote. Amastigotes of T. cruzi replicate inside the host cell; thus, it is difficult to conduct a knockout experiment without going through other developmental stages. Recently, our group reported a growth condition in which the amastigote can replicate axenically for up to 10 days without losing its amastigote-like properties. By using this temporal axenic amastigote culture, we successfully introduced gRNAs directly into the Cas9-expressing amastigote to cause gene knockouts and analyzed their phenotypes exclusively in the amastigote stage. In this report, we describe a detailed protocol to produce in vitro derived extracellular amastigotes, and to utilize the axenic culture in a CRISPR/Cas9-mediated knockout experiment. The growth phenotype of knockout amastigotes can be evaluated either by cell counts of the axenic culture, or by replication of intracellular amastigote after host cell invasion. This method bypasses the parasite stage differentiation normally involved in producing a transgenic or a knockout amastigote. Utilization of the temporal axenic amastigote culture has the potential to expand the experimental freedom of stage-specific studies in T. cruzi.

Introducing a Gene Knockout Directly Into the Amastigote Stage of Trypanosoma cruzi Using the CRISPR/Cas9 System

Here, we describe a protocol to introduce a gene knockout into the extracellular amastigote of Trypanosoma cruzi, using the CRISPR/Cas9 system. The growth phenotype can be followed up either by cell counting of axenic amastigote culture or by proliferation of intracellular amastigotes after host cell invasion.

Trypanosoma cruzi is a pathogenic protozoan parasite that causes Chagas&#8217; disease mainly in Latin America. In order to identify a novel drug target ...

Cell Surface Receptor Identification Using Genome-Scale CRISPR/Cas9 Genetic Screens

Summary

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Chapters in this video

Using a Fluorescent PCR-capillary Gel Electrophoresis Technique to Genotype CRISPR/Cas9-mediated Knockout Mutants in a High-throughput Format

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

Primordial Germ Cell Transplantation for CRISPR/Cas9-based Leapfrogging in Xenopus

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

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

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

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

Pooled CRISPR-Based Genetic Screens in Mammalian Cells

Introducing a Gene Knockout Directly Into the Amastigote Stage of Trypanosoma cruzi Using the CRISPR/Cas9 System