Genome-wide CRISPR-Cas9 knockout screens aim to elucidate the relationship between genotype and phenotype by ablating gene expression on a genome-wide scale and studying the resulting phenotypic alterations. The approach utilises the CRISPR-Cas9 gene editing system, coupled with libraries of single guide RNAs (sgRNAs), which are designed to target every gene in the genome. Over recent years, the genome-wide CRISPR screen has emerged as a powerful tool for performing large-scale loss-of-function screens, with low noise, high knockout efficiency and minimal off-target effects.
Early studies in Caenorhabditis elegans [1] and Drosophila melanogaster [2] [3] saw large-scale, systematic loss of function (LOF) screens performed through saturation mutagenesis, demonstrating the potential of this approach to characterise genetic pathways and identify genes with unique and essential functions. The saturation mutagenesis technique was later applied in other organisms, for example zebrafish [4] [5] and mice. [6] [7]
Targeted approaches for gene knockdown emerged in the 1980s with techniques such as homologous recombination, [8] [9] trans-cleaving ribozymes, [10] [11] and antisense technologies. [12] [13]
By the year 2000, RNA interference (RNAi) technology had emerged as a fast, simple, and inexpensive technique for targeted gene knockdown, and was routinely being used to study in vivo gene function in C. elegans. [14] [15] [16] [17] Indeed, in the span of only a few years following its discovery by Fire et al. (1998), [18] almost all of the ~19,000 genes in C. elegans had been analysed using RNAi-based knockdown. [19]
The production of RNAi libraries facilitated the application of this technology on a genome-wide scale, and RNAi-based methods became the predominant approach for genome-wide knockdown screens.[ citation needed ]
Nevertheless, RNAi-based approaches to genome-wide knockdown screens have their limitations. For one, the high off-target effects cause issues with false-positive observations. [20] [21] Additionally, because RNAi reduces gene expression at the post-transcriptional level by targeting RNA, RNAi-based screens only result in partial and short-term suppression of genes. Whilst partial knockdown may be desirable in certain situations, a technology with improved targeting efficiency and fewer off-target effects was needed.[ citation needed ]
Since initial identification as a prokaryotic adaptive immune system, [22] the bacterial type II clustered regularly interspaced short palindrome repeats (CRISPR)/Cas9 system has become a simple and efficient tool for generating targeted LOF mutations. [23] It has been successfully applied to edit human genomes, and has started to displace RNAi as the dominant tool in mammalian studies. [24] In the context of genome-wide knockout screens, recent studies have demonstrated that CRISPR/Cas9 screens are able to achieve highly efficient and complete protein depletion, and overcome the off-target issues seen with RNAi screens. [25] [26] In summary, the recent emergence of CRISPR-Cas9 has dramatically increased our ability to perform large-scale LOF screens. The versatility and programmability of Cas9, coupled with the low noise, high knockout efficiency and minimal off-target effects, have made CRISPR the platform of choice for many researchers engaging in gene targeting and editing. [24] [27]
The clustered regularly interspaced short palindrome repeats (CRISPR)/Cas9 system is a gene-editing technology that can introduce double-strand breaks (DSBs) at a target genomic locus. By using a single guide RNA (sgRNA), the endonuclease Cas9 can be delivered to a specific DNA sequence where it cleaves the nucleotide chain. [28] The specificity of the sgRNA is determined by a 20-nt sequence, homologous to the genomic locus of interest, and the binding to Cas9 is mediated by a constant scaffold region of the sgRNA. The desired target site must be immediately followed (5’ to 3’) by a conserved 3 nucleotide protospacer adjacent motif (PAM). [29] [30] In order to repair the DSBs, the cell may use the highly error prone non-homologous end joining, or homologous recombination. By designing suitable sgRNAs, planned insertions or deletions can be introduced into the genome. In the context of genome-wide LOF screens, the aim is to cause gene disruption and knockout.[ citation needed ]
To perform CRISPR knockouts on a genome-wide scale, collections of sgRNAs known as sgRNA libraries, or CRISPR knockout libraries, must be generated. The first step in creating a sgRNA library is to identify genomic regions of interest based on known sgRNA targeting rules. [31] For example, sgRNAs are most efficient when targeting the coding regions of genes and not the 5’ and 3’ UTRs. Conserved exons present as attractive targets, and position relative to the transcription start site should be considered. [31] Secondly, all the possible PAM sites are identified and selected for. [31] On- and off-target activity should be analysed, as should GC content, and homopolymer stretches should be avoided. [31] The most commonly used Cas9 endonuclease, derived from Streptococcus pyogenes , recognises a PAM sequence of NGG. [32]
Furthermore, specific nucleotides appear to be favoured at specific locations. Guanine is strongly favoured over cytosine on position 20 right next to the PAM motif, and on position 16 cytosine is preferred over guanine. [33] For the variable nucleotide in the NGG PAM motif, it has been shown that cytosine is preferred and thymine disfavoured. [33] With such criteria taken into account, the sgRNA library is computationally designed around the selected PAM sites. [31] [33] [34]
Multiple sgRNAs (at least 4–6) should be created against every single gene to limit false-positive detection, and negative control sgRNAs with no known targets should be included. [31] [33] The sgRNAs are then created by in situ synthesis, amplified by PCR, and cloned into a vector delivery system.
Developing a new sgRNA library is a laborious and time-consuming process. In practice, researchers may select an existing library depending on their experimental purpose and cell lines of interest. As of February 2020, the most widely used resources for genome-wide CRISPR knockout screens have been the two Genome-Scale CRISPR Knock-Out (GeCKO) libraries created by the Zhang lab. [35] Available through Addgene, these lentiviral libraries respectively target human and mouse exons, and both are available as a one-vector system (where the sgRNAs and Cas9 are present on the same plasmid) or as a two-vector system (where the sgRNAs and Cas9 are present on separate plasmids). Each library is delivered as two half-libraries, allowing researchers to screen with 3 or 6 sgRNAs/gene. [36]
Aside from GeCKO, a number of other CRISPR libraries have been generated and made available through Addgene. The Sabatini & Lander labs currently have 7 separate human and mouse libraries, including targeted sublibraries for distinct subpools such as kinases and ribosomal genes (Addgene #51043–51048). Further, improvements to the specificity of sgRNAs have resulted in ‘second generation’ libraries, such as the Brie (Addgene #73632) and Brunello (Addgene #73178) libraries generated by the Doench and Root labs, and the Toronto knockout (TKO) library (Addgene #1000000069) generated by the Moffat lab. [36]
Targeted gene knockout using CRISPR/Cas9 requires the use of a delivery system to introduce the sgRNA and Cas9 into the cell. Although a number of different delivery systems are potentially available for CRISPR, [37] [38] genome-wide loss-of-function screens are predominantly carried out using third generation lentiviral vectors. [35] [39] [40] These lentiviral vectors are able to efficiently transduce a broad range of cell types and stably integrate into the genome of dividing and non-dividing cells. [41] [42] Third generation lentiviral particles are produced by co-transfecting 293T human embryonic kidney (HEK) cells with:
The lentiviral particle-containing supernatant is harvested, concentrated and subsequently used to infect the target cells. [45] The exact protocol for lentiviral production will vary depending on the research aim and applied library. [35] [43] [44] If a two vector-system is used, for example, cells are sequentially transduced with Cas9 and sgRNA in a two-step procedure. [35] [44] Although more complex, this has the advantage of a higher titre for the sgRNA library virus. [35]
In general, there are two different formats of genome-wide CRISPR knockout screens: arrayed and pooled. In an arrayed screen, each well contains a specific and known sgRNA targeting a specific gene. [46] Since the sgRNA responsible for each phenotype is known based on well location, phenotypes can be identified and analysed without requiring genetic sequencing. This format allows for the measurement of more specific cellular phenotypes, perhaps by fluorescence or luminescence, and allows researchers to use more library types and delivery methods. [46] For large-scale LOF screens, however, arrayed formats are considered low-efficiency, and expensive in terms of financial and material resources because cell populations have to be isolated and cultured individually. [46]
In a pooled screen, cells grown in a single vessel are transduced in bulk with viral vectors collectively containing the entire sgRNA library. To ensure that the amount of cells infected by more than one sgRNA-containing particle is limited, a low multiplicity of infection (MOI) (typically 0.3-0.6) is used. [46] [47] Evidence so far has suggested that each sgRNA should be represented in a minimum of 200 cells. [48] [23] Transduced cells will be selected for, followed by positive or negative selection for the phenotype of interest, and genetic sequencing will be necessary to identify the integrated sgRNAs. [46]
Following phenotypic selection, genomic DNA is extracted from the selected clones, alongside a control cell population. [23] [46] [49] In the most common protocols for genome-wide knockouts, a 'Next-generation sequencing (NGS) library' is created by a two step polymerase chain reaction (PCR). [23] [46] The first step amplifies the sgRNA region, using primers specific to the lentiviral integration sequence, and the second step adds Illumina i5 and i7 sequences. [23] NGS of the PCR products allows the recovered sgRNAs to be identified, and a quantification step can be used to determine the relative abundance of each sgRNA. [23]
The final step in the screen is to computationally evaluate the significantly enriched or depleted sgRNAs, trace them back to their corresponding genes, and in turn determine which genes and pathways could be responsible for the observed phenotype. Several algorithms are currently available for this purpose, with the most popular being the Model-based Analysis of Genome-wide CRISPR/Cas9 Knockout (MAGeCK) method. [50] Developed specifically for CRISPR/Cas9 knockout screens in 2014, MAGeCK demonstrated better performance compared with alternative algorithms at the time, [50] and has since demonstrated robust results and high sensitivity across different experimental conditions. [51] As of 2015, the MAGeCK algorithm has been extended to introduce quality control measurements, and account for the previously overlooked sgRNA knockout efficiency. [51] A web-based visualisation tool (VISPR) was also integrated, allowing users to interactively explore the results, analysis, and quality controls. [51]
Over recent years, the genome-wide CRISPR screen has emerged as a powerful tool for studying the intricate networks of cellular signaling. [52] Cellular signaling is essential for a number of fundamental biological processes, including cell growth, proliferation, differentiation, and apoptosis.
One practical example is the identification of genes required for proliferative signaling in cancer cells. Cells are transduced with a CRISPR sgRNA library, and studied for growth over time. By comparing sgRNA abundance in selected cells to a control, one can identify which sgRNAs become depleted and in turn which genes may be responsible for the proliferation defect. Such screens have been used to identify cancer-essential genes in acute myeloid leukemia [53] and neuroblastoma, [54] and to describe tumor-specific differences between cancer cell lines. [55]
Targeted cancer therapies are designed to target the specific genes, proteins, or environments contributing to tumor cell growth or survival. After a period of prolonged treatment with these therapies, however, tumor cells may develop resistance. Although the mechanisms behind cancer drug resistance are poorly understood, potential causes include: target alteration, drug degradation, apoptosis escape, and epigenetic alterations. [56] Resistance is well-recognised and poses a serious problem in cancer management.[ citation needed ]
To overcome this problem, a synthetic lethal partner can be identified. Genome-wide LOF screens using CRISPR-Cas9 can be used to screen for synthetic lethal partners. [57] For this, a wild-type cell line and a tumor cell line containing the resistance-causing mutation are transduced with a CRISPR sgRNA library. The two cell lines are cultivated, and any under-represented or dead cells are analyzed to identify potential synthetic lethal partner genes. A recent study by Hinze et al. (2019) [58] used this method to identify a synthetic lethal interaction between the chemotherapy drug asparaginase and two genes in the Wnt signalling pathway NKD2 and LGR6.
Due to their small genomes and limited number of encoded proteins, viruses exploit host proteins for entry, replication, and transmission. Identification of such host proteins, also termed host dependency factors (HDFs), is particularly important for identifying therapeutic targets. Over recent years, many groups have successfully used genome-wide CRISPR/Cas9 as a screening strategy for HDFs in viral infections. [59]
One example is provided by Marceau et al. (2017), [60] who aimed to dissect the host factors associated with dengue and hepatitis C (HCV) infection (two viruses in family Flaviviridae). ELAVL1, an RNA-binding protein encoded by the ELAVL1 gene, was found to be a critical receptor for HCV entry, and a remarkable divergence in host dependency factors was demonstrated between the two flaviviridae. [60]
Additional reported applications of genome-wide CRISPR screens include the study of: mitochondrial metabolism, [61] bacterial toxin resistance, [62] genetic drivers of metastasis, [63] cancer drug resistance, [64] West Nile virus-induced cell death, [65] and immune cell gene networks. [66] [67]
This section will specifically address genome-wide CRISPR screens. For a review of CRISPR limitations see Lino et al. (2018) [38]
Genome-wide CRISPR screens will ultimately be limited by the properties of the chosen sgRNA library. Each library will contain a different set of sgRNAs, and average coverage per gene may vary. Currently available libraries tend to be biased towards sgRNAs targeting early (5’) protein-coding exons, rather than those targeting the more functional protein domains. [58] This problem was highlighted by Hinze et al. (2019), [58] who noted that genes associated with asparaginase sensitivity failed to score in their genome-wide screen of asparaginase-resistant leukemia cells.
If an appropriate library is not available, creating and amplifying a new sgRNA library is a lengthy process which may take many months. Potential challenges include: (i) effective sgRNA design; (ii) ensuring comprehensive sgRNA coverage throughout the genome; (iii) lentiviral vector backbone design; (iv) producing sufficient amounts of high-quality lentivirus; (v) overcoming low transformation efficiency; (vi) proper scaling of the bacterial culture. [68]
One of the largest hurdles for genome-wide CRISPR screening is ensuring adequate coverage of the sgRNA library across the cell population. [23] Evidence so far has suggested that each sgRNA should be represented and maintained in a minimum of 200-300 cells. [23] [48]
Considering that the standard protocol uses a multiplicity of infection of ~0.3, and a transduction efficiency of 30-40% [44] [23] the number of cells required to produce and maintain suitable coverage becomes very large. By way of example, the most popular human sgRNA library is the GeCKO v2 library created by the Zhang lab; [30] it contains 123,411 sgRNAs. Studies using this library commonly transduce more than 1x108 cells [58] [59] [69]
As CRISPR continues to exhibit low noise and minimal off-target effects, an alternative strategy is to reduce the number of sgRNAs per gene for a primary screen. Less stringent cut-offs are used for hit selection, and additional sgRNAs are later used in a more specific secondary screen. This approach is demonstrated by Doench et al. (2016), [33] who found that >92% of genes recovered using the standard protocol were also recovered using fewer sgRNAs per gene. They suggest that this strategy could be useful in studies where scale-up is prohibitively costly.[ citation needed ]
Lentiviral vectors have certain general limitations. For one, it is impossible to control where the viral genome integrates into the host genome, and this may affect important functions of the cell. Vannucci et al. [70] provide an excellent review of viral vectors along with their general advantages and disadvantages. In the specific context of genome-wide CRISPR screens, producing and transducing the lentiviral particles is relatively laborious and time-consuming, taking about two weeks in total. [44] Additionally, because the DNA integrates into the host genome, lentiviral delivery leads to long-term expression of Cas9, potentially leading to off-target effects.[ citation needed ]
In an arrayed screen, each well contains a specific and known sgRNA targeting a specific gene. Arrayed screens therefore allow for detailed profiling of a single cell, but are limited by high costs and the labour required to isolate and culture the high number of individual cell populations. [46] Conventional pooled CRISPR screens are relatively simple and cost effective to perform, but are limited to the study of the entire cell population. This means that rare phenotypes may be more difficult to identify, and only crude phenotypes can be selected for e.g. cell survival, proliferation, or reporter gene expression.[ citation needed ]
The choice of culture medium might affect the physiological relevance of findings from cell culture experiments due to the differences in the nutrient composition and concentrations. [71] A systematic bias in generated datasets was recently shown for CRISPR and RNAi gene silencing screens (especially for metabolic genes), [72] and for metabolic profiling of cancer cell lines. [71] For example, a stronger dependence on ASNS (asparagine synthetase) was found in cell lines cultured in DMEM, which lacks asparagine, compared to cell lines cultured in RPMI or F12 (containing asparagine). [72] Avoiding such bias might be achieved by using a uniform media for all screened cell lines, and ideally, using a growth medium that better represents the physiological levels of nutrients. Recently, such media types, as Plasmax [73] and Human Plasma Like Medium (HPLM), [74] were developed.
Emerging technologies are aiming to combine pooled CRISPR screens with the detailed resolution of massively parallel single-cell RNA-sequencing (RNA-seq). Studies utilising “CRISP-seq”, [75] “CROP-seq”, [76] and “PERTURB-seq” [77] have demonstrated rich genomic readouts, accurately identifying gene expression signatures for individual gene knockouts in a complex pool of cells. These methods have the added benefit of producing transcriptional profiles of the sgRNA-induced cells. [78]
Gene knockouts are a widely used genetic engineering technique that involves the targeted removal or inactivation of a specific gene within an organism's genome. This can be done through a variety of methods, including homologous recombination, CRISPR-Cas9, and TALENs.
A genetic screen or mutagenesis screen is an experimental technique used to identify and select individuals who possess a phenotype of interest in a mutagenized population. Hence a genetic screen is a type of phenotypic screen. Genetic screens can provide important information on gene function as well as the molecular events that underlie a biological process or pathway. While genome projects have identified an extensive inventory of genes in many different organisms, genetic screens can provide valuable insight as to how those genes function.
Gene knockdown is an experimental technique by which the expression of one or more of an organism's genes is reduced. The reduction can occur either through genetic modification or by treatment with a reagent such as a short DNA or RNA oligonucleotide that has a sequence complementary to either gene or an mRNA transcript.
Functional genomics is a field of molecular biology that attempts to describe gene functions and interactions. Functional genomics make use of the vast data generated by genomic and transcriptomic projects. Functional genomics focuses on the dynamic aspects such as gene transcription, translation, regulation of gene expression and protein–protein interactions, as opposed to the static aspects of the genomic information such as DNA sequence or structures. A key characteristic of functional genomics studies is their genome-wide approach to these questions, generally involving high-throughput methods rather than a more traditional "candidate-gene" approach.
CRISPR is a family of DNA sequences found in the genomes of prokaryotic organisms such as bacteria and archaea. Each sequence within an individual prokaryotic cell is derived from a DNA fragment of a bacteriophage that had previously infected the prokaryote or one of its ancestors. These sequences are used to detect and destroy DNA from similar bacteriophages during subsequent infections. Hence these sequences play a key role in the antiviral defense system of prokaryotes and provide a form of heritable, acquired immunity. CRISPR is found in approximately 50% of sequenced bacterial genomes and nearly 90% of sequenced archaea.
Guide RNA (gRNA) or single guide RNA (sgRNA) is a short sequence of RNA that functions as a guide for the Cas9-endonuclease or other Cas-proteins that cut the double-stranded DNA and thereby can be used for gene editing. In bacteria and archaea, gRNAs are a part of the CRISPR-Cas system that serves as an adaptive immune defense that protects the organism from viruses. Here the short gRNAs serve as detectors of foreign DNA and direct the Cas-enzymes that degrades the foreign nucleic acid.
Genome editing, or genome engineering, or gene editing, is a type of genetic engineering in which DNA is inserted, deleted, modified or replaced in the genome of a living organism. Unlike early genetic engineering techniques that randomly inserts genetic material into a host genome, genome editing targets the insertions to site-specific locations. The basic mechanism involved in genetic manipulations through programmable nucleases is the recognition of target genomic loci and binding of effector DNA-binding domain (DBD), double-strand breaks (DSBs) in target DNA by the restriction endonucleases, and the repair of DSBs through homology-directed recombination (HDR) or non-homologous end joining (NHEJ).
Cas9 is a 160 kilodalton protein which plays a vital role in the immunological defense of certain bacteria against DNA viruses and plasmids, and is heavily utilized in genetic engineering applications. Its main function is to cut DNA and thereby alter a cell's genome. The CRISPR-Cas9 genome editing technique was a significant contributor to the Nobel Prize in Chemistry in 2020 being awarded to Emmanuelle Charpentier and Jennifer Doudna.
CRISPR interference (CRISPRi) is a genetic perturbation technique that allows for sequence-specific repression of gene expression in prokaryotic and eukaryotic cells. It was first developed by Stanley Qi and colleagues in the laboratories of Wendell Lim, Adam Arkin, Jonathan Weissman, and Jennifer Doudna. Sequence-specific activation of gene expression refers to CRISPR activation (CRISPRa).
Epigenome editing or epigenome engineering is a type of genetic engineering in which the epigenome is modified at specific sites using engineered molecules targeted to those sites. Whereas gene editing involves changing the actual DNA sequence itself, epigenetic editing involves modifying and presenting DNA sequences to proteins and other DNA binding factors that influence DNA function. By "editing” epigenomic features in this manner, researchers can determine the exact biological role of an epigenetic modification at the site in question.
A protospacer adjacent motif (PAM) is a 2–6-base pair DNA sequence immediately following the DNA sequence targeted by the Cas9 nuclease in the CRISPR bacterial adaptive immune system. The PAM is a component of the invading virus or plasmid, but is not found in the bacterial host genome and hence is not a component of the bacterial CRISPR locus. Cas9 will not successfully bind to or cleave the target DNA sequence if it is not followed by the PAM sequence. PAM is an essential targeting component which distinguishes bacterial self from non-self DNA, thereby preventing the CRISPR locus from being targeted and destroyed by the CRISPR-associated nuclease.
CRISPR-Cas design tools are computer software platforms and bioinformatics tools used to facilitate the design of guide RNAs (gRNAs) for use with the CRISPR/Cas gene editing system.
Cas12a is an RNA-guided endonuclease that forms an essential component of the CRISPR systems found in some bacteria and archaea. In its natural context, Cas12a targets and destroys the genetic material of viruses and other foreign mobile genetic elements, thereby protecting the host cell from infection. Like other Cas enzymes, Cas12a binds to a "guide" RNA which targets it to a DNA sequence in a specific and programmable matter. In the host organism, the crRNA contains a constant region that is recognized by the Cas12a protein and a "spacer" region that is complementary to a piece of foreign nucleic acid that previously infected the cell.
No-SCAR genome editing is an editing method that is able to manipulate the Escherichia coli genome. The system relies on recombineering whereby DNA sequences are combined and manipulated through homologous recombination. No-SCAR is able to manipulate the E. coli genome without the use of the chromosomal markers detailed in previous recombineering methods. Instead, the λ-Red recombination system facilitates donor DNA integration while Cas9 cleaves double-stranded DNA to counter-select against wild-type cells. Although λ-Red and Cas9 genome editing are widely used technologies, the no-SCAR method is novel in combining the two functions; this technique is able to establish point mutations, gene deletions, and short sequence insertions in several genomic loci with increased efficiency and time sensitivity.
Perturb-seq refers to a high-throughput method of performing single cell RNA sequencing (scRNA-seq) on pooled genetic perturbation screens. Perturb-seq combines multiplexed CRISPR mediated gene inactivations with single cell RNA sequencing to assess comprehensive gene expression phenotypes for each perturbation. Inferring a gene’s function by applying genetic perturbations to knock down or knock out a gene and studying the resulting phenotype is known as reverse genetics. Perturb-seq is a reverse genetics approach that allows for the investigation of phenotypes at the level of the transcriptome, to elucidate gene functions in many cells, in a massively parallel fashion.
CRISPR activation (CRISPRa) is a gene regulation technique that utilizes an engineered form of the CRISPR-Cas9 system to enhance the expression of specific genes without altering the underlying DNA sequence. Unlike traditional CRISPR-Cas9, which introduces double-strand breaks to edit genes, CRISPRa employs a modified, catalytically inactive Cas9 (dCas9) fused with transcriptional activators to target promoter or enhancer regions, thereby boosting gene transcription. This method allows for precise control of gene expression, making it a valuable tool for studying gene function, creating gene regulatory networks, and developing potential therapeutic interventions for a variety of diseases.
Off-target genome editing refers to nonspecific and unintended genetic modifications that can arise through the use of engineered nuclease technologies such as: clustered, regularly interspaced, short palindromic repeats (CRISPR)-Cas9, transcription activator-like effector nucleases (TALEN), meganucleases, and zinc finger nucleases (ZFN). These tools use different mechanisms to bind a predetermined sequence of DNA (“target”), which they cleave, creating a double-stranded chromosomal break (DSB) that summons the cell's DNA repair mechanisms and leads to site-specific modifications. If these complexes do not bind at the target, often a result of homologous sequences and/or mismatch tolerance, they will cleave off-target DSB and cause non-specific genetic modifications. Specifically, off-target effects consist of unintended point mutations, deletions, insertions inversions, and translocations.
CRISPR gene editing (CRISPR, pronounced "crisper", refers to "clustered regularly interspaced short palindromic repeats") is a genetic engineering technique in molecular biology by which the genomes of living organisms may be modified. It is based on a simplified version of the bacterial CRISPR-Cas9 antiviral defense system. By delivering the Cas9 nuclease complexed with a synthetic guide RNA (gRNA) into a cell, the cell's genome can be cut at a desired location, allowing existing genes to be removed and/or new ones added in vivo.
Prime editing is a 'search-and-replace' genome editing technology in molecular biology by which the genome of living organisms may be modified. The technology directly writes new genetic information into a targeted DNA site. It uses a fusion protein, consisting of a catalytically impaired Cas9 endonuclease fused to an engineered reverse transcriptase enzyme, and a prime editing guide RNA (pegRNA), capable of identifying the target site and providing the new genetic information to replace the target DNA nucleotides. It mediates targeted insertions, deletions, and base-to-base conversions without the need for double strand breaks (DSBs) or donor DNA templates.
Genome editing of synthetic target arrays for lineage tracing (GESTALT) is a method used to determine the developmental lineages of cells in multicellular systems. GESTALT involves introducing a small DNA barcode that contains regularly spaced CRISPR/Cas9 target sites into the genomes of progenitor cells. Alongside the barcode, Cas9 and sgRNA are introduced into the cells. Mutations in the barcode accumulate during the course of cell divisions and the unique combination of mutations in a cell's barcode can be determined by DNA or RNA sequencing to link it to a developmental lineage.
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