Materials and Reagents

1. Petri dish (VWR, catalog number: 25384-088)
2. 12-well clear flat bottom not treated multiwell cell culture plate (Falcon, catalog number: 351143)
3. Glass bottom microwell dishes, 35 mm Petri dish, 14 mm microwell, No. 1.5 coverglass (0.16-0.19 mm) (MatTek, catalog number: P35G-1.5-14-C)
4. Glass transfer pipet (Fisher, catalog number: 63A183-624)
5. Double transgenic Tg(sox17:Cre^ERT2); Tg(ubi:Zebrabow) zebrafish (available upon request from the Amack lab: amackj@upstate.edu)
   Note: The Tg(sox17:Cre^ERT2); Tg(ubi:Zebrabow) strain is maintained by selecting embryos to raise that have green fluorescent hearts (cmlc2:GFP expression is a marker for the sox17: Cre^ERT2 transgene) and bright ubiquitous RFP expression from the ubi:Zebrabow transgene.
6. (Z)-4-Hydroxytamoxifen (4-OHT) (Sigma, catalog number: H7904)
   Note: Reconstituted to a stock solution of 10 mM in 1% DMSO and stored at -20 °C in single-use aliquots.
7. 1% low-melting point (LMP) agarose (Invitrogen, catalog number: 15517-014) prepared in embryo medium and maintained at 50 °C
8. 1% agarose (VWR, catalog number: 0710-500G) prepared in embryo medium that will be used to coat the bottom of Petri dishes and 12-well plates
9. Dimethyl sulfoxide (DMSO) (VWR, catalog number: 0231-500mL)
10. Sodium chloride (NaCl) (Fisher, catalog number: BP358-212)
11. Potassium chloride (KCl) (J. T. Baker, catalog number: 3040-01)
12. Calcium chloride dihydrate (CaCl₂•2H₂O) (J. T. Baker, catalog number: 1332-01)
13. Magnesium sulfate heptahydrate (MgSO₄•7H₂O) (Fisher, catalog number: BP214-500)
14. Methylene blue (Fisher, catalog number: BP117-100)
15. Embryo medium (see Recipes)
    

Equipment

1. Tweezers (Electron Microscopy Sciences, catalog number: 0103-5-PS)
   Note: We use Dumont Tweezer style 5.
2. Incubator set at 28.5 °C for culturing zebrafish embryos (VWR, catalog number: 35960-056)
3. Stereomicroscope with epifluorescent light source (Zeiss, model: Stereo Discovery V12)
   Note: We use a Zeiss Stereo Discovery V12.
4. Confocal microscope (Nikon, model: Eclipse Ti)
   Note: We use a Perkin-Elmer UltraVIEW Vox spinning disc confocal system equipped with 488 nm and 561 nm solid-state lasers (for YFP and RFP excitation) mounted on a Nikon Eclipse Ti inverted microscope with a Hamamatsu C9100-50 EM-CCD camera. We use a 20x oil-immersion objective. The microscope is equipped with a temperature control chamber for live imaging. It is likely that this protocol can be adapted for use with a laser scanning confocal microscope, but we have not tested different microscope platforms. 
5. Freezer
   

Software

1. Volocity (PerkinElmer) for image acquisition
2. Imaris (BitPlane) for 3D rendering and morphometric analysis
   

Procedure

1. Fluorescent mosaic labeling of DFC/KV cells
   
   1. Set up crosses of homozygous double transgenic Tg(sox17:Cre^ERT2); Tg(ubi:Zebrabow) zebrafish in breeding tanks with dividers that separate males from females. Remove dividers at the desired time to allow fish to breed and synchronize embryo development. 
   2. Collect Tg(sox17:Cre^ERT2); Tg(ubi:Zebrabow) embryos and culture them in embryo medium in a Petri dish at 28.5 °C until they reach the dome stage of development ~4 h post-fertilization (hpf).
   3. Carefully remove embryos from their chorion using fine tweezers (we use Dumont Tweezer style 5 from Electron Microscopy Sciences) in a Petri dish coated with 1% agarose. The agarose prevents the yolk of embryo from sticking to the plastic surface of Petri dish. To coat the dish, pipet enough hot liquid 1% agarose to cover the bottom of the dish, and then allow it to cool and solidify.
   4. Transfer dechorionated embryos using a glass transfer pipet (fire polish the tip) to a 12-well flat bottom cell culture plate coated with 1% agarose. Replace the embryo medium with fresh embryo medium containing 5 μM 4-hydroxytamoxifen (abbreviated here as 4-OHT) and 0.1% DMSO (dimethyl sulfoxide). The DMSO aids in cell permeability and drug delivery. Treat control embryos with 0.1% DMSO alone. We recommend that each well contain 5-6 dechorionated embryos. 
   5. Incubate embryos in 5 μM 4-OHT medium from the dome stage (4 hpf) to the shield stage (6 hpf) at 28.5 °C. 
   6. At the shield stage, transfer treated embryos to fresh embryo medium without 4-OHT and gently swirl. Repeat this step 3 times with fresh embryo medium to wash out 4-OHT. 
   7. Return rinsed embryos to 28.5 °C to allow development to the desired stage for imaging labeled DFC/KV cells.
      Note: Results from our work (Dasgupta et al., 2018) indicate Cre activity is not spatially biased, but randomly labels cells throughout the KV. In addition, we found on average that imaging 12 embryos will result in ~20 anterior KV cells and ~20 posterior KV cells to analyze. 
   
   
2. Immobilization of mosaic labeled embryos for imaging using an inverted microscope
   
   1. At the desired stage of development, carefully transfer an embryo to a glass-bottom (MatTek) dish using a glass transfer pipet. To analyze DFC behaviors, embryos can be prepared at any stage during epiboly. To visualize KV morphogenesis, we prepare embryos between the 1-2 somite stages.
      Note: Accumulation of YFP expression is time-dependent following Cre activation (4-OHT treatments). Thus, YFP fluorescence is weak at early (epiboly) stages and brighter at later (somite) stages.
   2. After transferring the embryo to the MatTek dish, remove most of the embryo medium and then cover the embryo with liquid 1% low-melting point (LMP) agarose that was maintained at 50 °C. 
   3. While the agarose solidifies, use a stereomicroscope to orient the embryo so that the DFC/KV cells face the glass-bottom (Figure 4; Video 1). 
      
      
      Figure 4. Immobilization of mosaic labeled embryos for live imaging. Schematic representing embryo immobilization technique used for live imaging with an inverted confocal microscope. A live embryo is covered with liquid low-melting point (LMP) agarose in a MatTek dish and then positioned such that DFC/KV cells are close to the glass bottom. Once solidified, the agarose is covered by embryo medium.
      
      
   4. Once the agarose has solidified, and the embryo is immobilized, add embryo medium to the dish to cover the sample and prevent the sample from drying out.
      Note: It is recommended to repeat this process to mount 5+ embryos for screening to identify the embryo(s) with the degree of mosaic labeling that is appropriate for the designed experiment.
      
      
      
      Video 1. Immobilization of a live embryo for confocal microscopy. This video demonstrates Steps 1-4 of Procedure B.
      
      
3. Imaging mosaic labeled DFC/KV cells in live embryos
   
   1. Position the MatTek dish containing the immobilized live embryo on an inverted confocal microscope. We use a 20x objective on a Perkin-Elmer UltraVIEW Vox spinning confocal disc confocal system with an environmental chamber maintained at 32 °C to image live embryos.
      Note: The Tg(ubi:Zebrabow) transgene drives RFP expression in all cells by default, which we excite using a 561 nm laser. If Cre-mediated recombination has occurred in a cell, we observe YFP expression (excited with a 488 nm laser).
   2. Select an embryo with bright YFP⁺ mosaic labeling that allows individual cells to be distinguished from their neighbors (Figure 3D). Laser power and exposure time will depend on signal intensity. Typically, we use 488 nm laser power between 30% and 50%, and exposure times between 500 ms and 800 ms using the 20x objective. Power for the 561 nm laser is typically 30% with an exposure time of 100-300 ms.
      Note: Laser power and exposure time should be minimized to prevent photo-toxicity. To achieve this, we suggest selecting mosaic labeled embryos with the brightest YFP expression.
   3. To analyze a single time point, we capture a Z-series through the entire KV using 2 µm Z steps. The typical distance is ~35 Z steps (70 μm) at the 2 somite stage and ~45 Z steps (90 μm) at the 8 somite stage. 
   4. For time-lapse imaging, we capture Z-stacks through the entire KV every 5 min during KV morphogenesis.
      Note: We have imaged a single embryo (using 5 min intervals) for up to 3 h (between 2 ss and 8 ss) without detecting photo-damage or deleterious effects on embryo development. 
   
   
4. Viewing confocal images using Imaris software
   
   1. To open a confocal image (Z-stack) in Imaris software, the raw data will need to be converted to an OME TIFF (.ome) file. In Volocity software, select the image to convert, and then right-click to export the file. Save the file as an OME TIFF.
   2. In Imaris (we have used version 8.4.0), click the Assay icon to create a new project folder. Next, click the Group icon to create a new group within the assay. Finally, click the Image icon and use the open file window to add an image (Z-stack in .ome file format) to the group.
   3. Double click on the .ome file icon to open the image in Imaris. This will open in the Surpass view.
   4. Next, open the Image Properties window (CTRL-I). Set the desired color for each channel (e.g., green, red, blue, etc.).
   5. Open the Geometry tab within the Image Properties window. Set the voxel size using pixel dimensions and Z-step size used to capture the image. Pixel dimensions are measured manually for each objective using a micrometer. For example:
      

      Spinning disc confocal 20x objective:	X = 0.33 (1 pixel = 0.33 μm)
      	Y = 0.33
      	Z = step size (2 μm) used to acquire Z-stack

   6. Open the Adjustment window (CTRL-D) to adjust signal levels. 
   7. Click Store to save the processed image.
   
   
5. 3D rendering of mosaic labeled DFC/KV cells using Imaris software
   
   1. To 3D render an object (e.g., single cell), click Surface under the 3D view menu.
   2. To define the region of interest (ROI) for rendering, go to the create tab and check Segment only a region of interest box.
   3. Click the next (blue) button at the bottom of the menu sidebar. The ROI bounding box appears (Figure 5A).
   4. To change the size of the bounding box, the cursor must be in the Select mode. Use the ESC key to toggle between Navigate and Select modes.
   5. In Select mode, click and drag arrowheads to re-size the bounding box in X, Y and Z around a single cell that you would like to reconstruct (Figure 5B).
      Note: you cannot zoom or rotate in Select mode; you must toggle to Navigate mode.
   6. Click next.
   7. Select source signal (e.g., green channel that includes YFP⁺ cells).
   8. Set surface detail. A higher number is more smooth and less detail (e.g., 1 = smooth).
   9. Set thresholding. Fifteen micrometer works well for KV cells. Smaller background signals are ignored.
      Note: The thresholding number should be based on the size of the object (cell) that you are rendering. The length and width of the cell can be measured manually in the 2D slice view.
   10. Click next.
   11. The slider can be used to manually adjust level of 3D rendering of the cell. Use navigate mode to check the rendering in X, Y, and Z.
   12. Optional: If a cell of interest is in contact with another labeled cell, use Split Touching Objects function. Check Enable and set the Seed Point Diameter. A value of 8 is often good for KV cells.
   13. Click next.
   14. Dots show how many labeled cells the software detects. 
   15. If necessary, go back and change Seed Point Diameter until the number of cells is accurate.
   16. Click the next button to finish. This completes the 3D rendering a single KV cell completes (Figure 5C). 
   17. To edit a 3D rendering, select edit (pencil icon in the lower menu bar). 
   18. Select the cell you wish to analyze and click duplicate in the edit tab. This creates a new surface file with only the selected 3D rendered cell.
   19. To obtain measurements of the 3D rendering, click statistics (graph icon in the lower menu bar), select detailed tab, select specific values and use the drop-down menu to select a measurement (e.g., area, volume, intensity, etc.). 
   20. Use clipping plane (scissors icon) to slice through the 3D rendered image. This tool can be used to slice 3D surface rendered KV cells to measure cell cross-sectional areas. 
   21. Use snapshot to capture an image (3D rendering, cross-section, etc.). Under the file tab, make a copy and save.
   22. Representative data are shown in Figure 6.
       
       
       Figure 5. 3D rendering of a single cell using a region of interest in Imaris software. A. A region of interest (ROI) bounding box (yellow box) around an entire 3D image of mosaic-labeled KV cells. Scale bar = 30 μm. B. The ROI bounding box can be re-sized to include only a single cell. Scale bar = 20 μm. C. The software 3D renders only the cell included in the ROI. Scale bar = 30 μm.
       
       
       Figure 6. Quantification of 3D rendered KV cells using Imaris. A. 3D renderings of cells in the anterior or posterior region of KV at the 2 somite stage (2 ss) or 8 somite stage (8 ss). Cell height (h), length (l), and width (w) measurements are shown at 2 ss. Dashed line indicates KV lumen surface. Scale bars = 10 μm. B-C. Box and whisker plots showing quantification of length to width ratio (LWR) that describes the shape of KV anterior or posterior cells (B), and quantification of cell volumes (C). n = 27 anterior cells at 2 ss; n = 25 posterior cells at 2 ss; n = 21 anterior cells at 8 ss; n = 22 posterior cells at 8 ss. Anterior and posterior KV cells have similar shapes and volumes at 2 ss, and then undergo asymmetric morphological changes that result in different cell shapes and volumes at 8 ss. n = number of cells analyzed. NS = not significant; ****P < 0.001. These images and results are modified from Dasgupta et al. (2018). 

Data analysis

Taking measurements of 3D rendered DFC/KV cells using Imaris software.

1. Click new measurement points icon in the top menu bar with small icons.
2. In the settings tab, select sphere (check the box).
3. For line mode check pairs.
4. Select edit tab.
5. Under intersect with, select surface of an object.
6. In Select mode, a box appears at the cursor.
7. Hold down the shift key and click to mark the first point of the line.
8. Repeat to mark the second point of the line.
9. To adjust the line, select one point (turns yellow), hold down shift and drag to new location.
   
10. Use settings tab to change font or color of the line.
11. Click statistics icon to obtain measurements in the detailed tab. Results can be saved as an excel file by clicking the export statistics (floppy disc icon) at the bottom of the menu sidebar.
12. Use snapshot to capture an image. Under the file tab, make a copy and save.
    

Notes

We realized Imaris is expensive software that is not available at all institutions. Alternative software packages such as Volocity (Perkin Elmer) or FIJI/ImageJ (free download at https://imagej.net/Fiji/Downloads) can be used to visualize confocal data sets in 3D, and generate 3D reconstructions for analysis. We focus this protocol on using Imaris because of its capability to segment a region of interest and 3D reconstruct a single cell (Figure 5).

Recipes

1. Embryo medium
   
   

Acknowledgments

We thank Sharleen Buel for technical assistance and zebrafish care, Madeline Clark for video demonstration, and Peu Santra for assistance with data presentation. The Tg(ubi:Zebrabow-M)^a131 strain was a kind gift from the Alex Schier Lab (Harvard University). This protocol corresponds to experiments presented in (Dasgupta et al., 2018). This work was supported by NIH grants R01HL095690 and R01GM117598.

Competing interests

The authors have no conflicts of interest to declare.

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