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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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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 (70um)</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&#39;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 CO2 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 (1 x PBS)</td>
			<td></td>
			<td></td>
			<td>8 g NaCl, 0.2 g KCl, 1.44 g Na2HPO4, 0.24 g KH2HPO4, dH2O up to 1L</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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Research

JoVE Journal

Developmental Biology

9.0K 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

Method of Studying Palatal Fusion using Static Organ Culture

Cleft lip and palate are among the most common of all birth defects. The secondary palate forms from mesenchymal shelves covered with epithelium that adheres to form the midline epithelial seam (MES). The theories suggest that MES cells follow an epithelial to mesenchymal transition (EMT), apoptosis and migration, making a fused palate 1. Complete disintegration of the MES is the final essential phase of palatal confluence with surrounding mesenchymal cells. We provide a method for palate organ culture. The developed in vitro protocol allows the study of the biological and molecular processes during fusion. The applications of this technique are numerous, including evaluating responses to exogenous chemical agents, effects of regulatory and growth factors and specific proteins. Palatal organ culture has a number of advantages including manipulation at different stages of development that is not possible using in vivo studies.

Studies of palate development are motivated by the incidence of cleft palate, a birth defect that imposes a tremendous health burden and can leave lasting disfigurement. We demonstrate here a technique to culture palatal shelves that can be used to study different signaling pathways involved in palatal development and fusion.

Cleft lip and palate are among the most common of all birth defects. The secondary palate forms from mesenchymal shelves covered with epithelium that ...

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 ...

Microinjection for Transgenesis and Genome Editing in Threespine Sticklebacks

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and behavioral phenotypes. The remarkably diverse phenotypes that have evolved as marine populations adapt to countless freshwater environments, combined with the ability to cross marine and freshwater forms, provide a rare vertebrate system in which genetics can be used to map genomic regions controlling evolved traits. Excellent genomic resources are now available, facilitating molecular genetic dissection of evolved changes. While mapping experiments generate lists of interesting candidate genes, functional genetic manipulations are required to test the roles of these genes. Gene regulation can be studied with transgenic reporter plasmids and BACs integrated into the genome using the Tol2 transposase system. Functions of specific candidate genes and cis-regulatory elements can be assessed by inducing targeted mutations with TALEN and CRISPR/Cas9 genome editing reagents. All methods require introducing nucleic acids into fertilized one-cell stickleback embryos, a task made challenging by the thick chorion of stickleback embryos and the relatively small and thin blastomere. Here, a detailed protocol for microinjection of nucleic acids into stickleback embryos is described for transgenic and genome editing applications to study gene expression and function, as well as techniques to assess the success of transgenesis and recover stable lines.

Transgenic manipulations and genome editing are critical for functionally testing the roles of genes and cis-regulatory elements. Here a detailed microinjection protocol for the generation of genomic modifications (including Tol2-mediated fluorescent reporter transgene constructs, TALENs, and CRISPRs) is presented for the emergent model fish, the threespine stickleback.

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and ...

Generation of Parabiotic Zebrafish Embryos by Surgical Fusion of Developing Blastulae

Surgical parabiosis of two animals of different genetic backgrounds creates a unique scenario to study cell-intrinsic versus cell-extrinsic roles for candidate genes of interest, migratory behaviors of cells, and secreted signals in distinct genetic settings. Because parabiotic animals share a common circulation, any blood or blood-borne factor from one animal will be exchanged with its partner and vice versa. Thus, cells and molecular factors derived from one genetic background can be studied in the context of a second genetic background. Parabiosis of adult mice has been &#160;used extensively to research aging, cancer, diabetes, obesity, and brain development. More recently, parabiosis of zebrafish embryos has been used to study the developmental biology of hematopoiesis. In contrast to mice, the transparent nature of zebrafish embryos permits the direct visualization of cells in the parabiotic context, making it a uniquely powerful method for investigating fundamental cellular and molecular mechanisms. The utility of this technique, however, is limited by a steep learning curve for generating the parabiotic zebrafish embryos. This protocol provides a step-by-step method on how to surgically fuse the blastulae of two zebrafish embryos of different genetic backgrounds to investigate the role of candidate genes of interest. In addition, the parabiotic zebrafish embryos are tolerant to heat shock, making temporal control of gene expression possible. This method does not require a sophisticated set-up and has broad applications for studying cell migration, fate specification, and differentiation in vivo during embryonic development.

This protocol provides step-by-step instruction on how to generate parabiotic zebrafish embryos of different genetic backgrounds. When combined with the unparalleled imaging capabilities of the zebrafish embryo, this method provides a uniquely powerful means to investigate cell-autonomous versus non-cell-autonomous functions for candidate genes of interest.

Surgical parabiosis of two animals of different genetic backgrounds creates a unique scenario to study cell-intrinsic versus cell-extrinsic roles for ...

Cell Lineage Analyses and Gene Function Studies Using Twin-spot MARCM

Mosaic analysis with a repressible cell marker (MARCM) is a positive mosaic labeling system that has been widely applied in Drosophila neurobiological studies to depict intricate morphologies and to manipulate the function of genes in subsets of neurons within otherwise unmarked and unperturbed organisms. Genetic mosaics generated in the MARCM system are mediated through site-specific recombination between homologous chromosomes within dividing precursor cells to produce both marked (MARCM clones) and unmarked daughter cells during mitosis. An extension of the MARCM method, called twin-spot MARCM (tsMARCM), labels both of the twin cells derived from a common progenitor with two distinct colors. This technique was developed to enable the retrieval of useful information from both hemi-lineages. By comprehensively analyzing different pairs of tsMARCM clones, the tsMARCM system permits high-resolution neural lineage mapping to reveal the exact birth-order of the labeled neurons produced from common progenitor cells. Furthermore, the tsMARCM system also extends gene function studies by permitting the phenotypic analysis of identical neurons of different animals. Here, we describe how to apply the tsMARCM system to facilitate studies of neural development in Drosophila.

Here, we present a protocol for a mosaic labeling technique that permits the visualization of neurons derived from a common progenitor cell in two distinct colors. This facilitates neural lineage analysis with the capability of birth-dating individual neurons and studying gene function in the same neurons of different individuals.

Mosaic analysis with a repressible cell marker (MARCM) is a positive mosaic labeling system that has been widely applied in Drosophila neurobiological ...

Structure-function Studies in Mouse Embryonic Stem Cells Using Recombinase-mediated Cassette Exchange

Gene engineering in mouse embryos or embryonic stem cells (mESCs) allows for the study of the function of a given protein. Proteins are the workhorses of the cell and often consist of multiple functional domains, which can be influenced by posttranslational modifications. The depletion of the entire protein in conditional or constitutive knock-out (KO) mice does not take into account this functional diversity and regulation. An mESC line and a derived mouse model, in which a docking site for FLPe recombination-mediated cassette exchange (RMCE) was inserted within the ROSA26 (R26) locus, was previously reported. Here, we report on a structure-function approach that allows for molecular dissection of the different functionalities of a multidomain protein. To this end, RMCE-compatible mice must be crossed with KO mice and then RMCE-compatible KO mESCs must be isolated. Next, a panel of putative rescue constructs can be introduced into the R26 locus via RMCE targeting. The candidate rescue cDNAs can be easily inserted between RMCE sites of the targeting vector using recombination cloning. Next, KO mESCs are transfected with the targeting vector in combination with an FLPe recombinase expression plasmid. RMCE reactivates the promoter-less neomycin-resistance gene in the ROSA26 docking sites and allows for the selection of the correct targeting event. In this way, high targeting efficiencies close to 100% are obtained, allowing for insertion of multiple putative rescue constructs in a semi-high throughput manner. Finally, a multitude of R26-driven rescue constructs can be tested for their ability to rescue the phenotype that was observed in parental KO mESCs. We present a proof-of-principle structure-function study in p120 catenin (p120ctn) KO mESCs using endoderm differentiation in embryoid bodies (EBs) as the phenotypic readout. This approach enables the identification of important domains, putative downstream pathways, and disease-relevant point mutations that underlie KO phenotypes for a given protein.

Proteins often contain multiple domains that can exert different cellular functions. Gene knock-outs (KO) do not consider this functional diversity. Here, we report a recombination-mediated cassette exchange (RMCE)-based structure-function approach in KO embryonic stem cells that allows for the molecular dissection of various functional domains or variants of a protein.

Gene engineering in mouse embryos or embryonic stem cells (mESCs) allows for the study of the function of a given protein. Proteins are the workhorses of ...

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...

Derivation of Stem Cell Lines from Mouse Preimplantation Embryos

Mouse embryonic stem cell (mESC) derivation is the process by which pluripotent cell lines are established from preimplantation embryos. These lines retain the ability to either self-renew or differentiate under specific conditions. Due to these properties, mESC are a useful tool in regenerative medicine, disease modeling, and tissue engineering studies. This article describes a simple protocol to obtain mESC lines with high derivation efficiencies (60-80%) by culturing blastocysts from permissive mouse strains on feeder cells in defined medium supplemented with leukemia inhibitory factor. The protocol can also be applied to efficiently derive mESC lines from non-permissive mouse strains, by the simple addition of a cocktail of two small-molecule inhibitors to the derivation medium (2i medium). Detailed procedures on the preparation and culture of feeder cells, collection and culture of mouse embryos, and derivation and culture of mESC lines are provided. This protocol does not require specialized equipment and can be carried out in any laboratory with basic mammalian cell culture expertise.

This article describes a protocol to efficiently derive and culture pluripotent stem cell lines from mouse embryos at the blastocyst stage.

Mouse embryonic stem cell (mESC) derivation is the process by which pluripotent cell lines are established from preimplantation embryos. These lines ...

Live Cell Imaging of Chromosome Segregation During Mitosis

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled by multiple mechanisms that include the Spindle Assembly Checkpoint (SAC). The SAC is part of a complex feedback system that is responsible for prevention of a cell progress through mitosis unless all chromosomal kinetochores have attached to spindle microtubules. Chromosomal lagging and abnormal chromosome segregation is an indicator of dysfunctional cell cycle control checkpoints and can be used to measure the genomic stability of dividing cells. Deregulation of the SAC can result in the transformation of a normal cell into a malignant cell through the accumulation of errors during chromosomal segregation. Implementation of the SAC and the formation of the kinetochore complex are tightly regulated by interactions between kinases and phosphatase such as Protein Phosphatase 2A (PP2A). This protocol describes live cell imaging of lagging chromosomes in mouse embryonic fibroblasts isolated from mice that had a knockout of the PP2A-B56&#947; regulatory subunit. This method overcomes the shortcomings of other cell cycle control imaging techniques such as flow cytometry or immunocytochemistry that only provide a snapshot of a cell cytokinesis status, instead of a dynamic spatiotemporal visualization of chromosomes during mitosis.

This protocol describes an easy and convenient method to label and visualize live chromosomes in mitotic cells using Histone2B-GFP BacMam 2.0 label and a spinning disc confocal microscopy system.

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled ...

An Efficient Strategy for Generating Tissue-specific Binary Transcription Systems in Drosophila by Genome Editing

Binary transcription systems are powerful genetic tools widely used for visualizing and manipulating cell fate and gene expression in specific groups of cells or tissues in model organisms. These systems contain two components as separate transgenic lines. A driver line expresses a transcriptional activator under the control of tissue-specific promoters/enhancers, and a reporter/effector line harbors a target gene placed downstream to the binding site of the transcription activator. Animals harboring both components induce tissue-specific transactivation of a target gene expression. Precise spatiotemporal expression of the gene in targeted tissues is critical for unbiased interpretation of cell/gene activity. Therefore, developing a method for generating exclusive cell/tissue-specific driver lines is essential. Here we present a method to generate highly tissue-specific targeted expression system by employing a &#34;Clustered Regularly Interspaced Short Palindromic Repeat/CRISPR-associated&#34; (CRISPR/Cas)-based genome editing technique. In this method, the endonuclease Cas9 is targeted by two chimeric guide RNAs (gRNA) to specific sites in the first coding exon of a gene in the Drosophila genome to create double-strand breaks (DSB). Subsequently, using an exogenous donor plasmid containing the transactivator sequence, the cell-autonomous repair machinery enables homology-directed repair (HDR) of the DSB, resulting in precise deletion and replacement of the exon with the transactivator sequence. The knocked-in transactivator is expressed exclusively in cells where the cis-regulatory elements of the replaced gene are functional. The detailed step-by-step protocol presented here for generating a binary transcriptional driver expressed in Drosophila fgf/branchless-producing epithelial/neuronal cells can be adopted for any gene- or tissue-specific expression.

An Efficient Strategy for Generating Tissue-specific Binary Transcription Systems in Drosophila by Genome Editing

Here, we present a method for generating tissue-specific binary transcription systems in Drosophila by replacing the first coding exon of genes with transcription drivers. The CRISPR/Cas9-based method places a transactivator sequence under the endogenous regulation of a replaced gene, and consequently facilitates transctivator expression exclusively in gene-specific spatiotemporal patterns.

Binary transcription systems are powerful genetic tools widely used for visualizing and manipulating cell fate and gene expression in specific groups of ...