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JoVE Journal Developmental Biology

Developmental Biology

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

Published: December 7, 2019 doi: 10.3791/60550
Automatic Translation
1,2, 3
1Photonics Group, Department of Physics, Imperial College London, 2The Medical Research Council Cancer Unit, Hutchison/MRC Research Centre, Cambridge Biomedical Campus, University of Cambridge, 3Cambridge Institute for Medical Research, Cambridge Biomedical Campus, University of Cambridge

Summary

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.

Abstract

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.

Introduction

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.

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Protocol

1. Differential Fluorescent Cell Labelling

  1. Gene tagging with CRISPR Cas9
    1. 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.
  2. Fluorescent dye labelling
    1. Grow Cal51 human cancer cells in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum (FBS), L-glutamine, and 100 µg/mL penicillin/streptomycin at 37 °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 (>15 passages). Cells must be healthy and growing exponentially. Avoid under-confluency or over-confluency.
    2. 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 70−90%.
      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.
    3. The next day, prewarm DMEM, trypsin and 1x phosphate-buffered saline (PBS) by placing them in a water bath at 37 °C.
    4. Wash rootletin-meGFP and rootletin-mScarlet-I cells 2x in PBS by pouring or aspirating off medium and replacing with 10 mL of PBS.
    5. Label Cal51 rootletin-meGFP cells by adding 500 nM violet cell dye (Table of Materials) in PBS for 1 min at room temperature (RT).
    6. 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.
    7. Stop the dye labelling reactions by adding 10 mL of DMEM for 5 min.
    8. 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).
    9. Wash all cells once by pouring off growth medium and replacing with 10 mL of PBS.
    10. Pour off PBS and incubate for 5 min at culture conditions with 1 mL of pre-warmed 1x trypsin.
    11. Transfer cells to a 15 mL plastic conical tube and centrifuge at 1000 x g for 5 min.
    12. Carefully remove and discard trypsin from the cell pellet with a pipette.
    13. Gently resuspend the violet and far red labelled cell pellets in 1 mL of PBS.
    14. 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.

2. Cell-cell Fusion

  1. 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.
  2. 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.
  3. 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.
  4. 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.
  5. 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.
  6. Add 10 mL of serum-free DMEM dropwise for 30 s and leave in the incubator for 10 min.
  7. 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.

3. Fluorescence-activated Cell Sorting (FACS) to Enrich Fused Cells

  1. Prepare all cells for flow cytometry by gently pipetting each sample separately through a 70 µm 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.
  2. Use a cytometer capable of single cell sorting.
    1. Set up the cell sorter for an aseptic sort. Perform the instrument quality control as per the manufacturer's recommendation.
    2. Draw a plot of forward scatter versus side scatter and a doublet discrimination plot.
  3. Create gates to identify fused cells.
    1. 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.
    2. Run unlabeled parental cells through the cytometer and record the fluorescence intensity.
      NOTE: Values above this baseline define positivity for dye staining.
    3. Run single labelled violet cells through the cytometer. Confirm that there is no spill-over of violet into the far red channel.
    4. 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 µm nozzle.
    5. Briefly run the fusion sample to confirm that fused cells are visible with the gates created in step 3.3.
  4. Align the sorting stream centrally into an 8-well imaging dish.
  5. FACS sort fused cells, present as double positive violet and far red labelled cells directly into an 8-well imaging dish containing 100 µL 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 20−90% 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.
  6. Directly after sorting, take cells back to the incubator and leave for >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.

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.

  1. Live cell imaging
    1. Replace growth medium with imaging medium without phenol red (Table of Materials) and proceed directly to imaging.
  2. Fixation and staining
    1. Prepare fresh 4% paraformaldehyde (PFA) in PBS.
    2. Fix cells by incubating them in 100 µL 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 °C protected from light if required.
    3. Wash cells 3x in 200 µL of PBS at RT.
    4. Permeabilize cells for 10 min at RT in 200 µL of 0.1% nonionic surfactant (Table of Materials) and 0.1% nonionic detergent (Table of Materials) diluted in PBS.
    5. Block in 200 µL of 3% bovine serum albumin in PBS for 30 min at RT.
    6. Incubate with antibodies in 150 µL 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.
    7. Wash 2x for 5 min in 300 µL of PBS and then leave in 200 µL of PBS.
      NOTE: Samples are imaged in PBS. Samples can be stored at 4 °C protected from light.
  3. Image acquisition
    1. Acquire images with an appropriate fluorescence microscope capable of four-color imaging (e.g., confocal, widefield, structured illumination).
    2. Excite meGFP and mScarlet-I channels with 488 nm and 561 nm wavelength lasers, respectively. Set detectors to detect at ~505−550 nm and ~590−650 nm. Excite violet and far red dye channels with 405 nm and 633 nm lasers, respectively. Set detectors to detect at ~430−500 nm and ~660−750 nm.
    3. Empirically determine laser intensity such that significant photobleaching does not occur and that cells remain healthy (if live cell imaging).
    4. Empirically determine gain such that signal is obtained without saturating or artificially clipping pixel values.
    5. Collect Z-stack data, such that images are three-dimensional, covering a range of 20−60 µm with ~500 µm 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.

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Representative Results

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.

Fusion induces major rearrangement of cellular architecture through mixing of two cells into one (Figure 2B). Heterokaryons are identified on the microscope as cells containing both fluorescent dye signals mixed inside a single cell (without intervening plasma membranes). Additionally, two nuclei may be visible in fused cells by brightfield/differential interference contrast or fluorescence imaging. Note that triploid or other more highly polyploid states are possible in addition to diploid fusions however, and so dye signal of two colors should be used to confirm the identity of fused cells.

Cell structure and function are further investigated through the merging of cells containing meGFP and mScarlet-I tagged proteins. Fusion results in a doubling of the centrosome number inside heterokaryons resulting from the fusion of two cells. Thus, if cells with fluorescently labelled centrosomes are fused, at least four pericentriolar material foci are observed when the centrosome pericentriolar component NEDD1 is fluorescently tagged (NEDD1-mRuby3; Figure 2C). Fusion of cells with endogenously fluorescently tagged rootletin (rootletin-meGFP and rootletin-mScarlet-I) allows the cell of origin of each centrosome to be identified in a heterokaryon. Rootletin in centrosomal roots has limited diffusional turnover24, and is therefore present as distinctly colored fibers in heterokaryons imaged with fluorescence microscopy (Figure 2D).

Figure 1
Figure 1: Cell-cell fusion and fluorescence imaging experimental workflow. Schematic of the four-stage experimental workflow. (1) Two cell populations are differentially labelled, with dyes and fluorescent fusion proteins. Cyan represents staining with violet cell dye and magenta represents staining with far red cell dye. Green represents meGFP tagging and red represents mScarlet-I tagging. (2) Cells are fused through incubation with polyethylene glycol. (3) Fused cells are enriched by flow cytometry, sorting the double fluorescently positive cells (far red and violet). (4) Fused cells are imaged by fluorescence microscopy to understand how cellular structure and function are altered (imaging the green and red channels). Please click here to view a larger version of this figure.

Figure 2
Figure 2: Representative flow cytometry enrichment and fluorescent imaging of fused cells. (A) Representative gating strategy used in flow cytometry sorting of fused cells. Fused cells that are doubly fluorescent are indicated by the black square. (B) Representative confocal fluorescence microscopy of fused cells, double labelled with violet and far red dyes. Shown are examples of successfully fused cells, which are either tetraploid or hexaploid (top and bottom panels respectively). Scale bar = 10 µm. (C) Representative live cell Airyscan confocal imaging of centrosomes in a single fused cell containing endogenously labelled centrosomal roots (rootletin-meGFP) and centrosomal pericentriolar material (NEDD1-mRuby3). Scale bar = 1 µm. (D) Cells expressing endogenously tagged rootletin-meGFP were fused with cells expressing endogenously tagged rootletin-mScarlet-I. Cells were fixed and stained and imaged by structured illumination microscopy. Shown is a maximum-intensity z-projection of centrosomes in one fused cell. Scale bar = 1 µm. Panel D has been modified from Mahen24 with permission. Please click here to view a larger version of this figure.

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Discussion

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. For example, when used at higher concentrations or with longer incubation times, cell dyes can be too bright and saturate the detectors during flow cytometry or fluorescence microscopy, or cause cross emission depending on imaging conditions. In the absence of a flow cytometer, other technologies are capable of enriching fused-cells, including double antibiotic selection27 and microfluidic trapping devices28. These technologies are either slower or require a more bespoke experimental setup, however.

Other methods of cell fusion have advantages and disadvantages in comparison to the protocol described here. Electro-fusion or viral-based fusion techniques may in some cases be imaged with microscopy during fusion8,29, making them good alternatives if it is preferable to observe the fusion process itself. However, these different methods may require specialized equipment (such as electrofusion equipment or viral transgenes). All cell-cell fusion methods have the potential to impinge upon cell health. Viral based fusion methods generally rely on the continued expression of viral transgenes, in contrast to the transient perturbation provided by PEG or electrofusion. Cellular fusogenic potential and toxicity after PEG exposure is variable in different cell types27, and hence titration of PEG incubation time may be required (protocol step 2.4), while recognizing that increased PEG exposure increases cell death30. Maximizing cell health is critical by keeping the time out of culture conditions to a minimum. Cell health can be checked during the protocol by measuring forward scatter versus side scatter at the cytometer and through observation of morphology during microscopy.

Careful consideration of experimental design to allow differential labelling with fluorescent tags is essential. The simplest design is using two separate fluorescent labels. However, endogenously expressed fluorescent fusion proteins are frequently of low expression level and hence are difficult to unambiguously detect by flow cytometry or imaging on the single cell level. We present a design that overcomes this limitation by including four fluorescent tags with CRISPR Cas9-mediated genome editing and dye-based staining methods. Fluorescent probes must have distinct emission spectra discernible from each other and be bright enough to distinguish from unlabeled cells on a single cell level. Probes must remain bound to a cell rather than dissipating externally. Irreversible binding internally on the subcellular level is not essential but may be desirable. Once cells merge, steady state exchange of cellular components can freely occur. Since roots are diffusionally stable, they allow tracing of cellular history, identifying the cell of origin of a given centrosome. A possible modification of the method is to tag other cellular structures of interest, but steady-state assemblies have the potential to mix after fusion, depending on the rate of intracellular dynamics (Figure 2C).

Cell-cell fusion induces a unique change in cellular architecture17,19,24, the implications of which are still not fully understood. This is both a limitation of the protocol and an exciting area for further investigation. Although it is clear that plasma membranes fuse into a single structure in heterokaryons31, how other cellular structures respond is poorly understood. Cell fusion could be used to further understand how organelle fusion and fission are regulated through the imaging of differentially labelled organelles24,32. Future work with cell fusion could address how euploidy, organelle number and cellular size impinges upon cell function. Since abnormal cell fusion has been observed in disease processes, in tumors33 and following viral infection10,17, this protocol is also of use for addressing a range of questions in fundamental and applied biology.

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Disclosures

The authors have nothing to disclose.

Acknowledgments

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.

Materials

Name Company Catalog Number Comments
15 mL tube Sarstedt 62554502
37% formaldehyde solution Sigma-Aldrich F8875
880 Laser Scanning Confocal Airyscan Microscope Carl Zeiss
8-well imaging dishes Ibidi 80826
Anti-GFP alpaca GFP booster nanobody Chromotek gba-488
BD Influx Cell Sorter BD Biosciences
Bovine serum albumin Sigma-Aldrich A7906
Cell Filters (70 μm) Biofil CSS010070
CellTrace Far Red ThermoFisher Scientific C34572
CellTrace Violet ThermoFisher Scientific C34571
Dulbecco's Modified Eagle Medium (DMEM), high glucose, GlutaMAX, pyruvate ThermoFisher Scientific 31966021
Fetal Bovine Serum Sigma-Aldrich 10270-106
FluoTag-X2 anti-mScarlet-I alpaca nanobody NanoTag Biotechnologies N1302-At565
L15 CO2 independent imaging medium Sigma-Aldrich 21083027
Penicillin/streptomycin Sigma-Aldrich 15140122
Phenol red free DMEM, high glucose ThermoFisher Scientific 21063029
Phosphate buffered saline (1x PBS) 8 g NaCl, 0.2 g KCl, 1.44 g Na2HPO4, 0.24 g KH2HPO4, dH2O up to 1 L
Polyethylene Glycol Hybri-Max 1450 Sigma-Aldrich P7181
Polypropylene tubes BD Falcon 352063
Triton X-100 Fisher BioReagents BP151 nonionic surfactant
Trypsin Sigma-Aldrich T4049
Tween 20 Fisher BioReagents BP337 nonionic detergent

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References

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Tags

Cell-cell Fusion Genome Editing Perturbation Cellular Structure Cellular Function Organelle Number Fluorescently Tagged Proteins Organelle Homeostasis Fluorescent Dye Labeling Cancer Cell Populations Confluency Cell Labeling Cell Cultures Trypsin Centrifugation
Cell-cell Fusion of Genome Edited Cell Lines for Perturbation of Cellular Structure and Function
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Mahen, R., Schulte, R. Cell-cell… More

Mahen, R., Schulte, R. Cell-cell Fusion of Genome Edited Cell Lines for Perturbation of Cellular Structure and Function. J. Vis. Exp. (154), e60550, doi:10.3791/60550 (2019).

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