Name: cell surface receptor identification using genome-scale crispr/cas9 genetic screens
Uploaded: 2020-06-06T13:30:00.000+00:00
Duration: 8 min 49 s
Description: Watch this Scientific Journal Video about Cell Surface Receptor Identification Using Genome-Scale CRISPR/Cas9 Genetic Screens at JoVE.com

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><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 &amp;amp; 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&amp;times; 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 ...

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Universidad de Cartagena UDEC

Research

JoVE Journal

Genetics

14.4K 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 ...

Investigation of Genetic Dependencies Using CRISPR-Cas9-based Competition Assays

Gene perturbation studies have been extensively used to investigate the role of individual genes in AML pathogenesis. For achieving complete gene disruption, many of these studies have made use of complex gene knockout models. While these studies with knockout mice offer an elegant and time-tested system for investigating genotype-to-phenotype relationships, a rapid and scalable method for assessing candidate genes that play a role in AML cell proliferation or survival in AML models will help accelerate the parallel interrogation of multiple candidate genes. Recent advances in genome-editing technologies have dramatically enhanced our ability to perform genetic perturbations at an unprecedented scale. One such system of genome editing is the CRISPR-Cas9-based method that can be used to make rapid and efficacious alterations in the target cell genome. The ease and scalability of CRISPR/Cas9-mediated gene-deletion makes it one of the most attractive techniques for the interrogation of a large number of genes in phenotypic assays. Here, we present a simple assay using CRISPR/Cas9 mediated gene-disruption combined with high-throughput flow-cytometry-based competition assays to investigate the role of genes that may play an important role in the proliferation or survival of human and murine AML cell lines.

This manuscript describes a Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) CRISPR-Cas9-based method for simple and expeditious investigation of the role of multiple candidate genes in Acute Myeloid Leukemia (AML) cell proliferation in parallel. This technique is scalable and can be applied in other cancer cell lines as well.

Gene perturbation studies have been extensively used to investigate the role of individual genes in AML pathogenesis. For achieving complete gene ...

Cancer Research

HOX Loci Focused CRISPR/sgRNA Library Screening Identifying Critical CTCF Boundaries

CCCTC-binding factor (CTCF)-mediated stable topologically associating domains (TADs) play a critical role in constraining interactions of DNA elements that are located in neighboring TADs. CTCF plays an important role in regulating the spatial and temporal expression of HOX genes that control embryonic development, body patterning, hematopoiesis, and leukemogenesis. However, it remains largely unknown whether and how HOX loci associated CTCF boundaries regulate chromatin organization and HOX gene expression. In the current protocol, a specific sgRNA pooled library targeting all CTCF binding sites in the HOXA/B/C/D loci has been generated to examine the effects of disrupting CTCF-associated chromatin boundaries on TAD formation and HOX gene expression. Through CRISPR-Cas9 genetic screening, the CTCF binding site located between HOXA7/HOXA9 genes (CBS7/9) has been identified as a critical regulator of oncogenic chromatin domain, as well as being important for maintaining ectopic HOX gene expression patterns in MLL-rearranged acute myeloid leukemia (AML). Thus, this sgRNA library screening approach provides novel insights into CTCF mediated genome organization in specific gene loci and also provides a basis for the functional characterization of the annotated genetic regulatory elements, both coding and noncoding, during normal biological processes in the post-human genome project era.

HOX Loci Focused CRISPR/sgRNA Library Screening Identifying Critical CTCF Boundaries

A CRISPR/sgRNA library has been applied to interrogating protein-coding genes.However, the feasibility of a sgRNA library to uncover the function of a CTCF boundary in gene regulation remains unexplored. Here, we describe a HOX loci specific sgRNA library to elucidate the function of CTCF boundaries in HOX loci.

CCCTC-binding factor (CTCF)-mediated stable topologically associating domains (TADs) play a critical role in constraining interactions of DNA elements ...

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 ...

CRISPR/Cas9 Editing of the C. elegans rbm-3.2 Gene using the dpy-10 Co-CRISPR Screening Marker and Assembled Ribonucleoprotein Complexes.

The bacterial Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR)/Streptococcus pyogenes CRISPR-associated protein (Cas) system has been harnessed by researchers to study important biologically relevant problems. The unparalleled power of the CRISPR/Cas genome editing method allows researchers to precisely edit any locus of their choosing, thereby facilitating an increased understanding of gene function. Several methods for editing the C. elegans genome by CRISPR/Cas9 have been described previously. Here, we discuss and demonstrate a method which utilizes in vitro assembled ribonucleoprotein complexes and the dpy-10 co-CRISPR marker for screening. Specifically, in this article, we go through the step-by-step process of introducing premature stop codons into the C. elegans rbm-3.2 gene by homology-directed repair using this method of CRISPR/Cas9 editing. This relatively simple editing method can be used to study the function of any gene of interest and allows for the generation of homozygous-edited C. elegans by CRISPR/Cas9 editing&#160;in less than two weeks.

CRISPR/Cas9 Editing of the C. elegans rbm-3.2 Gene using the dpy-10 Co-CRISPR Screening Marker and Assembled Ribonucleoprotein Complexes.

The goal of this protocol is to enable seamless CRISPR/Cas9 editing of the C. elegans genome using assembled ribonucleoprotein complexes and the dpy-10 co-CRISPR marker for screening. This protocol can be used to make a variety of genetic modifications in C. elegans including insertions, deletions, gene replacements and codon substitutions.

The bacterial Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR)/Streptococcus pyogenes CRISPR-associated protein (Cas) system has been ...

Detecting Off-Target CRISPR-Cas9 Cleavage Events with CIRCLE-Seq

CIRCLE-Seq for Interrogation of Off-Target Gene Editing

Circularization for In Vitro Reporting of Cleavage Effects by Sequencing (CIRCLE-seq) is a novel technique developed for the impartial identification of unintended cleavage sites of CRISPR-Cas9 through targeted sequencing of CRISPR-Cas9 cleaved DNA. The protocol involves circularizing genomic DNA (gDNA), which is subsequently treated with the Cas9 protein and a guide RNA (gRNA) of interest. Following treatment, the cleaved DNA is purified and prepared as a library for Illumina sequencing. The sequencing process generates paired-end reads, offering comprehensive data on each cleavage site. CIRCLE-seq provides several advantages over other in vitro methods, including minimal sequencing depth requirements, low background, and high enrichment for Cas9-cleaved gDNA. These advantages enhance sensitivity in identifying both intended and unintended cleavage events. This study provides a comprehensive, step-by-step procedure for examining the off-target activity of CRISPR-Cas9 using CIRCLE-seq. As an example, this protocol is validated by mapping genome-wide unintended cleavage sites of CRISPR-Cas9 during the modification of the AAVS1 locus. The entire CIRCLE-seq process can be completed in two weeks, allowing sufficient time for cell growth, DNA purification, library preparation, and Illumina sequencing. The input of sequencing data into the CIRCLE-seq pipeline facilitates streamlined interpretation and analysis of cleavage sites.

A significant barrier to technologies like CRISPR is the off-target events that can disrupt vital genes. 'Circularization for In Vitro Reporting of Cleavage Effects by Sequencing' (CIRCLE-seq) is a technique designed to identify unintended cleavage sites. This method maps the genome-wide activity of CRISPR-Cas9 with high sensitivity and without bias.

Circularization for In Vitro Reporting of Cleavage Effects by Sequencing (CIRCLE-seq) is a novel technique developed for the impartial identification of ...

InSET: A Novel Tool for Unbiased Genome-Wide Functional Element Discovery

Identification of Functionally-Relevant Lentivirus Integration Sites in an Insertional Mutagenesis Cell Library

The extent of functional sequences within the human genome is a pivotal yet debated topic in biology. Although high-throughput reverse genetic screens have made strides in exploring this, they often limit their scope to known genomic elements and may introduce non-specific effects. This underscores the urgent need for novel functional genomics tools that enable a deeper, unbiased understanding of genome functionality. This protocol introduces the Insertion-based Screen for functional Elements and Transcripts (InSET), a method for identifying lentivirus integration sites within a lentivirus-based insertional mutagenesis cell library. InSET facilitates the capture of genome-wide lentiviral integration sites, with next-generation sequencing used to detect and quantify flanking sequences. InSET's design enables the analysis of integration site abundance variations in phenotypic screens on a large scale, establishing it as a robust tool for forward genetics and for identifying functional genomic elements. A key benefit of InSET is its capacity to reveal previously unidentified genomic elements, including novel functional exons of both protein-coding and non-coding RNAs, independent of prior annotation. Overall, InSET holds significant value in studying the intricate complexity of the human genome and transcriptome, where many genomic elements await functional characterization.

Insertional mutagenesis is an essential tool in forward genetics for identifying functional genomic elements. Here, we describe the Insertion-based Screen for functional Elements and Transcripts (InSET), a method for detecting lentivirus integration sites within a lentivirus-based insertional mutagenesis cell library.

The extent of functional sequences within the human genome is a pivotal yet debated topic in biology. Although high-throughput reverse genetic screens ...

Evaluation of Abnormal Growth-related Genes of Hematopoietic Stem and Progenitor Cells by Combining CRISPR/Cas9 Technology with Cell Counting

Hematopoietic stem cells possess the ability for long-term self-renewal and the potential to differentiate into various types of mature blood cells. However, the accumulation of cancerous mutations in hematopoietic stem and progenitor cells (HSPCs) can block normal differentiation, induce aberrant proliferation, and ultimately lead to leukemogenesis. To identify and/or evaluate the cancerous mutations, we integrated CRISPR/Cas9 technology with the Cell Counting Kit-8 (CCK-8) assay to investigate a model gene Trp53, that is essential for abnormal proliferative ability of HSPCs. Specifically, bone marrow cells enriched for HSPCs from Cas9 mice were harvested and then subjected to viral transfection with single-guide RNAs (sgRNAs) targeting one or several candidate genes to introduce genetic alterations in HSPCs. Then, a CCK-8 assay was performed to investigate the proliferative capacity of transfected HSPCs. The sgRNA targeting efficiency was confirmed by a Tracking of Indels by Decomposition assay. These identified genes may play crucial roles in leukemogenesis and could serve as potential therapeutic targets.

Here, we present a protocol that integrates CRISPR/Cas9 technology with the Cell Counting Kit-8 (CCK-8) assay to identify candidate genes that are crucial for the abnormal proliferative capacity of hematopoietic stem and progenitor cells.

Hematopoietic stem cells possess the ability for long-term self-renewal and the potential to differentiate into various types of mature blood cells. ...

In Vitro Selection of Engineered Transcriptional Repressors for Targeted Epigenetic Silencing

Gene inactivation is instrumental to study gene function and represents a promising strategy for the treatment of a broad range of diseases. Among traditional technologies, RNA interference suffers from partial target abrogation and the requirement for life-long treatments. In contrast, artificial nucleases can impose stable gene inactivation through induction of a DNA double strand break (DSB), but recent studies are questioning the safety of this approach. Targeted epigenetic editing via engineered transcriptional repressors (ETRs) may represent a solution, as a single administration of specific ETR combinations can lead to durable silencing without inducing DNA breaks.
ETRs are proteins containing a programmable DNA-binding domain (DBD) and effectors from naturally occurring transcriptional repressors. Specifically, a combination of three ETRs equipped with the KRAB domain of human ZNF10, the catalytic domain of human DNMT3A and human DNMT3L, was shown to induce heritable repressive epigenetic states on the ETR-target gene. The hit-and-run nature of this platform, the lack of impact on the DNA sequence of the target, and the possibility to revert to the repressive state by DNA demethylation on demand, make epigenetic silencing a game-changing tool. A critical step is the identification of the proper ETRs&#39; position on the target gene to maximize on-target and minimize off-target silencing. Performing this step in the final ex vivo or in vivo preclinical setting can be cumbersome.
Taking the CRISPR/catalytically dead Cas9 system as a paradigmatic DBD for ETRs, this paper describes a protocol consisting of the in vitro screen of guide RNAs (gRNAs) coupled to the triple-ETR combination for efficient on-target silencing, followed by evaluation of the genome-wide specificity profile of top hits. This allows for reduction of the initial repertoire of candidate gRNAs to a short list of promising ones, whose complexity is suitable for their final&#160;evaluation in the therapeutically relevant setting of interest.

In Vitro Selection of Engineered Transcriptional Repressors for Targeted Epigenetic Silencing

Here, we present a protocol for the in vitro selection of engineered transcriptional repressors (ETRs) with high, long-term, stable, on-target silencing efficiency and low genome-wide, off-target activity. This workflow allows for reducing an initial, complex repertoire of candidate ETRs to a short list, suitable for further evaluation in therapeutically relevant settings.

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

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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

Investigation of Genetic Dependencies Using CRISPR-Cas9-based Competition Assays

HOX Loci Focused CRISPR/sgRNA Library Screening Identifying Critical CTCF Boundaries

Pooled CRISPR-Based Genetic Screens in Mammalian Cells

CRISPR/Cas9 Editing of the C. elegans rbm-3.2 Gene using the dpy-10 Co-CRISPR Screening Marker and Assembled Ribonucleoprotein Complexes.

CIRCLE-Seq for Interrogation of Off-Target Gene Editing

Identification of Functionally-Relevant Lentivirus Integration Sites in an Insertional Mutagenesis Cell Library

Evaluation of Abnormal Growth-related Genes of Hematopoietic Stem and Progenitor Cells by Combining CRISPR/Cas9 Technology with Cell Counting

In Vitro Selection of Engineered Transcriptional Repressors for Targeted Epigenetic Silencing