This work was funded by a Wellcome Trust Henry Wellcome Fellowship to R.M. (https://wellcome.ac.uk/grant number 100090/12/Z). The funder had no role in the study design, data collection and analysis, decision to publish, or preparation of the manuscript. We thank Ashok Venkitaraman and Paul French for critical advice and guidance on the project. We thank Chiara Cossetti and Gabriela Grondys-Kotarba in the Cambridge Institute for Medical Research Flow Cytometry facility for excellent support. We thank Liam Cassiday, Thomas Miller, and Gianmarco Contino for proofreading the manuscript.

1. Differential Fluorescent Cell Labelling
<ol>
	<li>Gene tagging with CRISPR Cas9
	<ol>
		<li>Use CRISPR Cas9 genome editing to tag rootletin (or other genes of interest) with the fluorescent proteins meGFP or mScarlet-I in human cancer cell lines. 
		NOTE: Detailed protocols for genome editing are covered elsewhere24,25,26.</li>
	</ol>
	</li>
	<li>Fluorescent dye labelling
	<ol>
		<li>Grow Cal51 human cancer cells in Dulbecco&#39;s modified Eagle&#39;s medium (DMEM) supplemented with 10% fetal bovine serum (FBS), L-glutamine, and 100 &#181;g/mL penicillin/streptomycin at 37 &#176;C and 5% CO2 in a humidified incubator. 
		NOTE: It is important to ensure that cells are regularly checked for the absence of mycoplasma contamination, and for the identity of the cell line. Do not use cells that have been extensively passaged in culture (&#62;15 passages). Cells must be healthy and growing exponentially. Avoid under-confluency or over-confluency.</li>
		<li>Seed each cell type (rootletin-meGFP, rootletin-mScarlet-I and parental untagged Cal51 cells) such that ~6 x 106 cells are present the next day for each sample, at a confluency of 70&#8722;90%. 
		NOTE: This is approximately one T75 tissue culture flask or one 10 cm dish per cell type. The parental untagged cells are required as a negative control later in the protocol. This protocol allows for the production of ~20,000 fused cells, but this number could be increased through scaling up the protocol with an increased number of batches.</li>
		<li>The next day, prewarm DMEM, trypsin and 1x phosphate-buffered saline (PBS) by placing them in a water bath at 37 &#176;C.</li>
		<li>Wash rootletin-meGFP and rootletin-mScarlet-I cells 2x in PBS by pouring or aspirating off medium and replacing with 10 mL of PBS.</li>
		<li>Label Cal51 rootletin-meGFP cells by adding 500 nM violet cell dye (Table of Materials) in PBS for 1 min at room temperature (RT).</li>
		<li>Label Cal51 rootletin-mScarlet-I cells by adding 200 nM far red cell dye (Table of Materials) in PBS for 1 min at RT. 
		NOTE: Keep samples shielded from light wherever possible after fluorescent staining, to avoid photobleaching of fluorescence.</li>
		<li>Stop the dye labelling reactions by adding 10 mL of DMEM for 5 min.</li>
		<li>Remove unlabeled parental Cal51 cells from the incubator (from step 1.2.2). 
		NOTE: These cells will be used later as unlabeled negative controls (step 3.3.2).</li>
		<li>Wash all cells once by pouring off growth medium and replacing with 10 mL of PBS.</li>
		<li>Pour off PBS and incubate for 5 min at culture conditions with 1 mL of pre-warmed 1x trypsin.</li>
		<li>Transfer cells to a 15 mL plastic conical tube and centrifuge at 1000 x g for 5 min.</li>
		<li>Carefully remove and discard trypsin from the cell pellet with a pipette.</li>
		<li>Gently resuspend the violet and far red labelled cell pellets in 1 mL of PBS.</li>
		<li>Gently resuspend the parental (non-fluorescent) cell pellet in 1 mL of DMEM and return to the incubator. 
		NOTE: This sample will not undergo cell-cell fusion but will be used later during flow cytometry.</li>
	</ol>
	</li>
</ol>
2. Cell-cell Fusion
<ol>
	<li>Mix 3 x 106 violet labelled cells with 3 x 106 far red labelled cells in a 15 mL tube by gently pipetting together 0.5 mL of each cell type using a 1 mL pipette.</li>
	<li>Centrifuge the remaining unmixed cells from step 2.1 at 1,000 x g for 5 min, then pour off PBS, resuspend the pellet in 1 mL of DMEM and return to the incubator. 
	NOTE: These remaining unmixed single-labelled samples will be used later as negative and compensation controls in section 3 of the protocol.</li>
	<li>Centrifuge the mixed cells from step 2.1 at 1,000 x g for 5 min and carefully aspirate PBS using a 1 mL pipette.</li>
	<li>Add 0.7 mL of 50% 1450 PEG solution in a dropwise fashion to the cell pellet (step 2.3) using a 1 mL pipette over a period of 30 s.</li>
	<li>Leave for 3.5 min at RT. 
	NOTE: Incubation time with PEG may be optimized depending on cell type, although note that longer incubation time increases cell toxicity.</li>
	<li>Add 10 mL of serum-free DMEM dropwise for 30 s and leave in the incubator for 10 min.</li>
	<li>Spin down at 1,000 x g for 5 min, discard supernatant, and resuspend gently in 1 mL of complete medium (DMEM containing FBS). 
	NOTE: Proceed directly to section 3. There is toxicity associated with PEG exposure followed by flow cytometry and imaging, so keep cells in culture conditions as much as possible by proceeding rapidly between steps.</li>
</ol>
3. Fluorescence-activated Cell Sorting (FACS) to Enrich Fused Cells
<ol>
	<li>Prepare all cells for flow cytometry by gently pipetting each sample separately through a 70 &#181;m filter into a FACS tube. 
	NOTE: Four samples are required: fused cells, single labelled far red cells, single labelled violet cells, unlabeled parental cells. Once removed from the sterile tissue culture hood, cells have the capacity to become infected, so minimize the exposure time of samples to a non-sterile environment from here onwards. Cells are sorted in complete DMEM medium.</li>
	<li>Use a cytometer capable of single cell sorting.
	<ol>
		<li>Set up the cell sorter for an aseptic sort. Perform the instrument quality control as per the manufacturer&#39;s recommendation.</li>
		<li>Draw a plot of forward scatter versus side scatter and a doublet discrimination plot.</li>
	</ol>
	</li>
	<li>Create gates to identify fused cells.
	<ol>
		<li>Excite the fluorescent dyes at wavelengths of 405 nm and 635 nm. Detect at 450 nm and 660 nm (violet and far-red wavelengths, respectively). Plot far red versus violet wavelengths.</li>
		<li>Run unlabeled parental cells through the cytometer and record the fluorescence intensity. 
		NOTE: Values above this baseline define positivity for dye staining.</li>
		<li>Run single labelled violet cells through the cytometer. Confirm that there is no spill-over of violet into the far red channel.</li>
		<li>Run single labelled far red cells through the cytometer and confirm that there is no spill-over of far red into the violet channel. 
		NOTE: For the representative data shown, cells were sorted at 20 psi on a sorter equipped with a 100 &#181;m nozzle.</li>
		<li>Briefly run the fusion sample to confirm that fused cells are visible with the gates created in step 3.3.</li>
	</ol>
	</li>
	<li>Align the sorting stream centrally into an 8-well imaging dish.</li>
	<li>FACS sort fused cells, present as double positive violet and far red labelled cells directly into an 8-well imaging dish containing 100 &#181;L of growth medium. 
	NOTE: The maximum recommended number of cells is ~50,000 per 8-well dish. Ensure cell health during sorting. Cell confluency can influence cell health so keep cells between 20&#8722;90% confluency post-sorting by sorting an appropriate number of cells for the imaging dish. Each sorted droplet contains approximately 3 nL of sheath fluid. Ensure that sheath fluid does not significantly dilute the growth medium during sorting.</li>
	<li>Directly after sorting, take cells back to the incubator and leave for &#62;2 h or overnight. 
	NOTE: Adherent cells will gradually re-adhere to the coverslip during this period, from sorting onwards, and hence their morphology will change over time if taken directly to the microscope.</li>
</ol>
4. Immunofluorescent Staining and Imaging of Cell-cell Fusions
NOTE: Fused cells can be imaged live or after fixation and further fluorescent staining (or both), depending on the experiments and measurements required.
<ol>
	<li>Live cell imaging
	<ol>
		<li>Replace growth medium with imaging medium without phenol red (Table of Materials) and proceed directly to imaging.</li>
	</ol>
	</li>
	<li>Fixation and staining
	<ol>
		<li>Prepare fresh 4% paraformaldehyde (PFA) in PBS.</li>
		<li>Fix cells by incubating them in 100 &#181;L of 4% PFA for 15 min at RT. Remove PFA after 15 min. 
		CAUTION: PFA is toxic when inhaled so this step should be performed in a fume hood with appropriate personal protective equipment. 
		NOTE: After fixation, the experiment can be paused and restarted later if required. Store the sample at 4 &#176;C protected from light if required.</li>
		<li>Wash cells 3x in 200 &#181;L of PBS at RT.</li>
		<li>Permeabilize cells for 10 min at RT in 200 &#181;L of 0.1% nonionic surfactant (Table of Materials) and 0.1% nonionic detergent (Table of Materials) diluted in PBS.</li>
		<li>Block in 200 &#181;L of 3% bovine serum albumin in PBS for 30 min at RT.</li>
		<li>Incubate with antibodies in 150 &#181;L of PBS containing 3% bovine serum albumin and 0.1% nonionic surfactant and 0.05% nonionic detergent for 1 h at RT. 
		NOTE: The primary antibodies used to generate the representative results are fluorescently conjugated anti-GFP nanobody (used at 1:400 dilution), and fluorescently conjugated anti-mScarlet-I nanobody (used at 1:500 dilution). These enhance the signal of the already fluorescent proteins.</li>
		<li>Wash 2x for 5 min in 300 &#181;L of PBS and then leave in 200 &#181;L of PBS. 
		NOTE: Samples are imaged in PBS. Samples can be stored at 4 &#176;C protected from light.</li>
	</ol>
	</li>
	<li>Image acquisition
	<ol>
		<li>Acquire images with an appropriate fluorescence microscope capable of four-color imaging (e.g., confocal, widefield, structured illumination).</li>
		<li>Excite meGFP and mScarlet-I channels with 488 nm and 561 nm wavelength lasers, respectively. Set detectors to detect at ~505&#8722;550 nm and ~590&#8722;650 nm. Excite violet and far red dye channels with 405 nm and 633 nm lasers, respectively. Set detectors to detect at ~430&#8722;500 nm and ~660&#8722;750 nm.</li>
		<li>Empirically determine laser intensity such that significant photobleaching does not occur and that cells remain healthy (if live cell imaging).</li>
		<li>Empirically determine gain such that signal is obtained without saturating or artificially clipping pixel values.</li>
		<li>Collect Z-stack data, such that images are three-dimensional, covering a range of 20&#8722;60 &#181;m with ~500 &#181;m step size. 
		NOTE: The images shown in representative data were acquired by either confocal (Figure 2B), Airyscan (Figure 2C) or structured illumination microscopy (Figure 2D). Each cell is a bona fide heterokaryon if it contains both violet and far red dyes overlapping in the cytoplasm.</li>
	</ol>
	</li>
</ol>

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</ol>

The authors have nothing to disclose.

We demonstrate a facile and cost-effective protocol for fusing cells and visualizing the subsequent architecture of cell hybrids with microscopy, taking approximately two days from start to finish. Critical parts of this protocol are the enrichment of fused cells by cell sorting (protocol section 3), and careful validation of fused cells by microscopy (protocol section 4). These sections ensure that fused cells are readily obtained and are bona fide heterokaryons. Concentrations and incubation times should be adhered to....

Homeostatic maintenance of cellular structure is critical to life. Cells have characteristic morphologies, sub-cellular organelle numbers, and internal biochemical composition. Understanding how these fundamental properties are generated and how they go awry during disease requires laboratory tools to perturb them.
Cell fusion is the merging of two or more separate cells. Cell fusion may have been critical to the emergence of eukaryotic life1. In the human body, cell fusion is relatively rare, occurring during restricted developmental circumstances and tissue types, such as during fertilization or the formation of muscle, bone and the placenta2. This protocol describes the induction of cell-cell fusion in tissue culture cell lines with differentially fluorescently labelled organelles, as a tool to understand the mechanisms controlling cell structure and function.
In vitro induced cell-cell fusion is central to the production of monoclonal antibodies3, an important tool for biological research and disease treatment. Cell fusion has also been used to ask many different fundamental cell biological questions about cell cycle dominance4, aneuploidy5,6, cellular reprogramming7,8, the repair of damaged neurons9, viral proliferation10, apoptosis11, tumorigenesis12, cytoskeletal dynamics13, and membrane fusion14,15. Laboratory based methods to induce cell-cell fusion16,17,18,19 induce lipid membrane coalescence through the physical merging of two bilayers into one. Cell fusion can be induced by electricity18, viral based methods17, thermoplasmonic heating20, transgene expression19, and chemicals including polyethylene glycol (PEG)16,21,22.
Centrosomes are microtubule organizing centers controlling cellular shape, motility, polarization, and division23. Centrosomal roots are fibrous structures extending from centrosomes containing the protein rootletin24 (encoded by the gene CROCC). We recently used cell-cell fusion to understand how centrosome position and number varies inside heterokaryons relative to parental cells24. The rationale behind the use of this method is to track the cell of origin of roots within a heterokaryon after fusion of differentially fluorescently tagged parental cells, and thus to image organelle fusion and fission. The fluorescently tagged proteins rootletin-meGFP or rootletin-mScarlet-I are created by genome editing in separate cell lines which are then fused by PEG-mediated cell fusion. We describe the use of cell dyes (Table of Materials) to identify fused cells by flow cytometry and subsequent fluorescence microscopy identification of centrosome cell of origin and morphology (Figure 1). This approach is a robust and unique method to study how major changes in cellular state including organelle number impinge upon cell homeostasis.

<table><tbody><tr><td>15 mL tube</td><td>Sarstedt</td><td>62554502</td><td></td></tr><tr><td>37% formaldehyde solution</td><td>Sigma-Aldrich</td><td>F8875</td><td></td></tr><tr><td>880 Laser Scanning Confocal Airyscan Microscope</td><td>Carl Zeiss</td><td></td><td></td></tr><tr><td>8-well imaging dishes</td><td>Ibidi</td><td>80826</td><td></td></tr><tr><td>Anti-GFP alpaca GFP booster nanobody</td><td>Chromotek</td><td>gba-488</td><td></td></tr><tr><td>BD Influx Cell Sorter</td><td>BD Biosciences</td><td></td><td></td></tr><tr><td>Bovine serum albumin</td><td>Sigma-Aldrich</td><td>A7906</td><td></td></tr><tr><td>Cell Filters (70 &amp;mu;m)</td><td>Biofil</td><td>CSS010070</td><td></td></tr><tr><td>CellTrace Far Red</td><td>ThermoFisher Scientific</td><td>C34572</td><td></td></tr><tr><td>CellTrace Violet</td><td>ThermoFisher Scientific</td><td>C34571</td><td></td></tr><tr><td>Dulbecco&#039;s Modified Eagle Medium (DMEM), high glucose, GlutaMAX, pyruvate</td><td>ThermoFisher Scientific</td><td>31966021</td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>Sigma-Aldrich</td><td>10270-106</td><td></td></tr><tr><td>FluoTag-X2 anti-mScarlet-I alpaca nanobody</td><td>NanoTag Biotechnologies</td><td>N1302-At565</td><td></td></tr><tr><td>L15 CO&lt;sub&gt;2&lt;/sub&gt; independent imaging medium</td><td>Sigma-Aldrich</td><td>21083027</td><td></td></tr><tr><td>Penicillin/streptomycin</td><td>Sigma-Aldrich</td><td>15140122</td><td></td></tr><tr><td>Phenol red free DMEM, high glucose</td><td>ThermoFisher Scientific</td><td>21063029</td><td></td></tr><tr><td>Phosphate buffered saline (1x PBS)</td><td></td><td></td><td>8 g NaCl, 0.2 g KCl, 1.44 g Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;, 0.24 g KH&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;, dH&lt;sub&gt;2&lt;/sub&gt;O up to 1 L</td></tr><tr><td>Polyethylene Glycol Hybri-Max 1450</td><td>Sigma-Aldrich</td><td>P7181</td><td></td></tr><tr><td>Polypropylene tubes</td><td>BD Falcon</td><td>352063</td><td></td></tr><tr><td>Triton X-100</td><td>Fisher BioReagents</td><td>BP151</td><td>nonionic surfactant</td></tr><tr><td>Trypsin</td><td>Sigma-Aldrich</td><td>T4049</td><td></td></tr><tr><td>Tween 20</td><td>Fisher BioReagents</td><td>BP337</td><td>nonionic detergent</td></tr></tbody></table>

cell-cell fusion of genome edited cell lines for perturbation of cellular structure and function

Life is spatially partitioned within lipid membranes to allow the isolated formation of distinct molecular states inside cells and organelles. Cell fusion is the merger of two or more cells to form a single cell. Here we provide a protocol for cell fusion of two different cell types. Fused hybrid cells are enriched by flow cytometry-based sorting, followed by fluorescence microscopy of hybrid cell structure and function. Fluorescently tagged proteins generated by genome editing are imaged inside fused cells, allowing cellular structures to be identified based on fluorescence emission and referenced back to the cell type of origin. This robust and general method can be applied to different cell types or organelles of interest, to understand cellular structure and function across a range of fundamental biological questions.

Appropriately labelled cells are visible during flow cytometry by fluorescence signal higher than unlabeled control cells (Figure 2A). Gates are set for sorting of double positive cells, enriching this population directly into imaging dishes for further microscopic analyses. Fused cells are detectable as distinct double fluorescently positive cells and constitute about ~1% of the population.
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Watch this Scientific Journal Video about Cell-cell Fusion of Genome Edited Cell Lines for Perturbation of Cellular Structure and Function at JoVE.com

Cell-cell Fusion of Genome Edited Cell Lines for Perturbation of Cellular Structure and Function

The purpose of this protocol is to fuse two different cell types to create hybrid cells. Fluorescence microscopy analysis of fused cells is used to track the cell of origin of cellular organelles. This assay can be used to explore how cellular structure and function respond to perturbation by cell fusion.

Life is spatially partitioned within lipid membranes to allow the isolated formation of distinct molecular states inside cells and organelles. Cell fusion ...

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JoVE Journal

Developmental Biology

9.1K Views.  Imperial College London. The purpose of this protocol is to fuse two different cell types to create hybrid cells. Fluorescence microscopy analysis of fused cells is used to track the cell of origin of cellular organelles. This assay can be used to explore how cellular structure and function respond to perturbation by cell fusion.

Video: Cell-cell Fusion of Genome Edited Cell Lines for Perturbation of Cellular Structure and Function

Genome Editing and Directed Differentiation of hPSCs for Interrogating Lineage Determinants in Human Pancreatic Development

Interrogating gene function in self-renewing or differentiating human pluripotent stem cells (hPSCs) offers a valuable platform towards understanding human development and dissecting disease mechanisms in a dish. To capitalize on this potential application requires efficient genome-editing tools to generate hPSC mutants in disease-associated genes, as well as in vitro hPSC differentiation protocols to produce disease-relevant cell types that closely recapitulate their in vivo counterparts. An efficient genome-editing platform for hPSCs named iCRISPR has been developed through the TALEN-mediated targeting of a Cas9 expression cassette in the AAVS1 locus. Here, the protocols for the generation of inducible Cas9 hPSC lines using cells cultured in a chemically defined medium and a feeder-free condition are described. Detailed procedures for using the iCRISPR system for gene knockout or precise genetic alterations in hPSCs, either through non-homologous end joining (NHEJ) or via precise nucleotide alterations using a homology-directed repair (HDR) template, respectively, are included. These technical procedures include descriptions of the design, production, and transfection of CRISPR guide RNAs (gRNAs); the measurement of the CRISPR mutation rate by T7E1 or RFLP assays; and the establishment and validation of clonal mutant lines. Finally, we chronicle procedures for hPSC differentiation into glucose-responsive pancreatic &#946;-like cells by mimicking in vivo pancreatic embryonic development. Combining iCRISPR technology with directed hPSC differentiation enables the systematic examination of gene function to further our understanding of pancreatic development and diabetes disease mechanisms.

Protocols to generate hPSC mutant lines using the iCRISPR platform and to differentiate hPSCs into glucose-responsive &#946;-like cells are described. Combining genome editing technology with hPSC-directed differentiation provides a powerful platform for the systematic analysis of the role of lineage determinants in human development and disease progression.

Interrogating gene function in self-renewing or differentiating human pluripotent stem cells (hPSCs) offers a valuable platform towards understanding ...

Genetics

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

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

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

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

Establishment of Genome-edited Human Pluripotent Stem Cell Lines: From Targeting to Isolation

Genome-editing of human pluripotent stem cells (hPSCs) provides a genetically controlled and clinically relevant platform from which to understand human development and investigate the pathophysiology of disease. By employing site-specific nucleases (SSNs) for genome editing, the rapid derivation of new hPSC lines harboring specific genetic alterations in an otherwise isogenic setting becomes possible. Zinc finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs) and clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9 are the most commonly used SSNs. All of these nucleases function by introducing a double stranded DNA break at a specified site, thereby promoting precise gene editing at a genomic locus. SSN-meditated genome editing exploits two of the cell&#39;s endogenous DNA repair mechanisms, non-homologous end joining (NHEJ) and homology directed repair (HDR), to either introduce insertion/deletion mutations or alter the genome using a homologous repair template at the site of the double stranded break. Electroporation of hPSCs is an efficient means of transfecting SSNs and repair templates that incorporate transgenes such as fluorescent reporters and antibiotic resistance cassettes. After electroporation, it is possible to isolate only those hPSCs that incorporated the repair construct by selecting for antibiotic resistance. Mechanically separating hPSC colonies and confirming proper integration at the target site through genotyping allows for the isolation of correctly targeted and genetically homogeneous cell lines. The validity of this protocol is demonstrated here by using all three SSN platforms to incorporate EGFP and a puromycin resistance construct into the AAVS1 safe harbor locus in human pluripotent&#160;stem cells.

Genome editing of human pluripotent stem cells (hPSCs) can be done quickly and efficiently. Presented here is a robust experimental procedure to genetically engineer hPSCs as exemplified by editing the AAVS1 safe harbor locus to express EGFP and introduce antibiotic resistance.

Genome-editing of human pluripotent stem cells (hPSCs) provides a genetically controlled and clinically relevant platform from which to understand human ...

Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has quickly become a commonplace amongst researchers pursuing a wide variety of biological questions. Users most often employ the Cas9 protein derived from Streptococcus pyogenes in a complex with an easily reprogrammed guide RNA (gRNA). These components are introduced into cells, and through a base pairing with a complementary region of the double-stranded DNA (dsDNA) genome, the enzyme cleaves both strands to generate a double-strand break (DSB). Subsequent repair leads to either random insertion or deletion events (indels) or the incorporation of experimenter-provided DNA at the site of the break.
The use of a purified single-guide RNA and Cas9 protein, preassembled to form an RNP and delivered directly to cells, is a potent approach for achieving highly efficient gene editing. RNP editing particularly enhances the rate of gene insertion, an outcome that is often challenging to achieve. Compared to the delivery via a plasmid, the shorter persistence of the Cas9 RNP within the cell leads to fewer off-target events. 
Despite its advantages, many casual users of CRISPR gene editing are less familiar with this technique. To lower the barrier to entry, we outline detailed protocols for implementing the RNP strategy in a range of contexts, highlighting its distinct benefits and diverse applications. We cover editing in two types of primary human cells, T cells and hematopoietic stem/progenitor cells (HSPCs). We also show how Cas9 RNP editing enables the facile genetic manipulation of entire organisms, including the classic model roundworm Caenorhabditis elegans and the more recently introduced model crustacean, Parhyale hawaiensis.

Utilizing a preassembled Cas9 ribonucleoprotein complex (RNP) is a powerful method for precise, efficient genome editing. Here, we highlight its utility across a broad range of cells and organisms, including primary human cells and both classic and emerging model organisms.

Site-specific eukaryotic genome editing with CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) systems has ...

Genome Editing in Mammalian Cell Lines using CRISPR-Cas

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

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

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

Introducing Point Mutations into Human Pluripotent Stem Cells Using Seamless Genome Editing

Custom designed endonucleases, such as RNA-guided Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)-Cas9, enable efficient genome editing in mammalian cells. Here we describe detailed procedures to seamlessly genome edit the hepatocyte nuclear factor 4 alpha (HNF4&#945;) locus as an example in human pluripotent stem cells. Combining a piggyBac-based donor plasmid and the CRISPR-Cas9 nickase mutant in a two-step genetic selection, we demonstrate correct and efficient targeting of the HNF4&#945; locus.

Here, we describe a detailed method for seamless gene editing in human pluripotent stem cells using a piggyBac-based donor plasmid and the Cas9 nickase mutant. Two point mutations were introduced into exon 8 of the hepatocyte nuclear factor 4 alpha (HNF4&#945;) locus in human embryonic stem cells (hESCs).

Custom designed endonucleases, such as RNA-guided Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)-Cas9, enable efficient genome editing ...

Zinc-finger Nuclease Enhanced Gene Targeting in Human Embryonic Stem Cells

One major limitation with current human embryonic stem cell (ESC) differentiation protocols is the generation of heterogeneous cell populations.&#160;These cultures contain the cells of interest, but are also contaminated with undifferentiated ESCs, non-neural derivatives and other neuronal subtypes.&#160; This limits their use in in vitro and in vivo applications, such as in vitro modeling for drug discovery or cell replacement therapy. To help overcome this, reporter cell lines, which offer a means to visualize, track and isolate cells of interest, can be engineered.&#160;However, to achieve this in human embryonic stem cells via conventional homologous recombination is extremely inefficient.&#160;This protocol describes targeting of the Pituitary homeobox 3 (PITX3) locus in human embryonic stem cells using custom designed zinc-finger nucleases, which introduce site-specific double-strand DNA breaks, together with a PITX3-EGFP-specific DNA donor vector. Following the generation of the PITX3 reporter cell line, it can then be differentiated using published protocols for use in studies such as in vitro Parkinson&#8217;s disease modeling or cell replacement therapy.

Reporter cell lines offer a means to visualize, track and isolate cells of interest from heterogeneous populations.&#160;However, gene-targeting using conventional homologous recombination in human embryonic stem cells is extremely inefficient. Herein, we describe targeting CNS midbrain specific transcription factor PITX3 locus with EGFP using zinc-finger nuclease enhanced homologous recombination.

One major limitation with current human embryonic stem cell (ESC) differentiation protocols is the generation of heterogeneous cell ...

Biology

Generation of Genomic Deletions in Mammalian Cell Lines via CRISPR/Cas9

The prokaryotic clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated (Cas) 9 system may be re-purposed for site-specific eukaryotic genome engineering. CRISPR/Cas9 is an inexpensive, facile, and efficient genome editing tool that allows genetic perturbation of genes and genetic elements. Here we present a simple methodology for CRISPR design, cloning, and delivery for the production of genomic deletions. In addition, we describe techniques for deletion, identification, and characterization. This strategy relies on cellular delivery of a pair of chimeric single guide RNAs (sgRNAs) to create two double strand breaks (DSBs) at a locus in order to delete the intervening DNA segment by non-homologous end joining (NHEJ) repair. Deletions have potential advantages as compared to single-site small indels given the efficiency of biallelic modification, ease of rapid identification by PCR, predictability of loss-of-function, and utility for the study of non-coding elements. This approach can be used for efficient loss-of-function studies of genes and genetic elements in mammalian cell lines.

CRISPR/Cas9 is a robust system to produce disruption of genes and genetic elements. Here we describe a protocol for the efficient creation of genomic deletions in mammalian cell lines using CRISPR/Cas9.

The prokaryotic clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated (Cas) 9 system may be re-purposed for site-specific ...

Genome Editing

A well-established technique for modifying specific sequences in the genome is gene targeting by homologous recombination, but this method can be laborious and only works in certain organisms. Recent advances have led to the development of &#8220;genome editing&#8221;, which works by inducing double-strand breaks in DNA using engineered nuclease enzymes guided to target genomic sites by either proteins or RNAs that recognize specific sequences. When a cell attempts to repair this damage, mutations can be introduced into the targeted DNA region. In this video, JoVE explains the principles behind genome editing, emphasizing how this technique relates to DNA repair mechanisms. Then, three major genome editing methods&#8212;zinc finger nucleases, TALENs, and the CRISPR-Cas9 system&#8212;are reviewed, followed by a protocol for using CRISPR to create targeted genetic changes in mammalian cells. Finally, we discuss some current research that applies genome editing to alter the genetic material in model organisms or cultured cells.

Genome Editing: ZFNs, TALENs, and CRISPR-cas9 System

A well-established technique for modifying specific sequences in the genome is gene targeting by homologous recombination, but this method can be ...

Education

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Advanced Biology

All-in-One CRISPR Genome Editing: A Method for Homology Directed Repair-Based Gene Knock-In in Cultured Cells Using CRISPR-Cas9 System

Source: Zhang, L. et al, <a href="https://www.jove.com/v/62328">Gene Knock-in by CRISPR/Cas9 and Cell Sorting in Macrophage and T Cell Lines.</a> J. Vis. Exp. (2021).
In this video, we demonstrate all-in-one CRISPR-Cas9 based genome editing in cultured cells where Cas9 and sgRNA are provided as a single plasmid construct to the cells. The CRISPR-Cas9 system and desired gene to be inserted was introduced in cells through electroporation technique to facilitate successful gene editing.

Source: Zhang, L. et al, Gene Knock-in by CRISPR/Cas9 and Cell Sorting in Macrophage and T Cell Lines. J. Vis. Exp. (2021).
In this video, we demonstrate ...