Live Imaging and Characterization of Microglia Dynamics in the Zebrafish Embryo

Zebrafish were maintained under standard conditions, according to FELASA42 and institutional (Université Libre de Bruxelles, Brussels, Belgium; ULB) guidelines and regulations. All experimental procedures were approved by the ULB Ethical Committee for Animal Welfare (CEBEA) from the ULB.

1. Zebrafish husbandry and embryo preparation

NOTE: The zebrafish transgenic line Tg(mpeg1:eGFP)^gl22 that expresses the fluorescent eGFP protein in macrophages, including microglia, was used to generate the tracking data shown in this protocol. Other macrophage reporter lines are available from ZIRC and can also be used. In choosing the transgenic line, it is important to consider that a high signal intensity will facilitate image acquisition and cell segmentation.

1. Maintain zebrafish (Danio rerio) at a temperature of 28 °C and manage them in accordance with established protocols¹¹.
2. Stimulate mating by placing female and male fish together in a breeding tank, with a mesh bottom and a separator, in the late afternoon before the desired embryos' birth date. Remove the separator the following morning to time the mating of the fish.
3. Collect the fertilized eggs using a mesh and transfer them to a 100 mm Petri dish containing E3 medium (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl₂, 0.33 mM MgSO₄) with 0.01% Methylene blue. Depending on the genetic background of the zebrafish transgenic line used, if pigment formation is to be inhibited to facilitate imaging, add 0.003% of N-phenylthiourea (PTU) to the E3 buffer in which embryos are developing, starting at 6 h post fertilization (hpf) and refreshing the solution every 2 days until the desired developmental stage is reached.
   NOTE: To avoid long exposure to PTU that could affect embryonic development¹², it is suggested to use, if available, a transgenic line on a pigment-deficient background, such as Casper^w2, that allows for clear imaging without the need to expose embryos to PTU for an extended period of time¹³.
4. If the embryos to be imaged are younger than 3 days post fertilization (dpf), dechorionate them at 24 hpf. Use a stereo microscope to confirm that all embryos have been dechorionated and are undamaged by the enzymatic treatment (see step 1.4.2).
   1. In case of a limited number of eggs, do this manually by using fine dissecting forceps to pinch the chorion and pull outwards, carefully releasing each embryo from its chorion.
   2. In case of a higher number of eggs, perform the dechorionation enzymatically by incubating the eggs in bulk at room temperature (RT) in a 1 mg/mL Pronase solution in E3 medium for a brief period (typically 5-10 min, until the chorions break open under gently swishing). Next, the Pronase solution is discarded, and the embryos are rinsed thoroughly 3x with fresh E3, to remove any remaining chorion and residual enzyme.

2. Zebrafish mounting

1. Prepare a 1% low melting agarose solution in E3 buffer. Cool the agarose to 37 °C in a benchtop heating block and add freshly made Tricaine methanesulfonate solution (160 mg/L), which will help keep the embryos anesthetized during imaging. Vortex the solution to thoroughly homogenize it.
2. Anesthetize the embryos by adding 160 mg/L of Tricaine methanesulfonate to the Petri dish containing the embryos in E3 buffer. Check to verify that movement has stopped and select the zebrafish embryos with the desired fluorescence. Check the embryo's health by visualizing its heartbeat using a brightfield stereo microscope. Using a wide-tipped transfer pipette draw up one, or up to three, of the selected embryos and transfer them to a bottom glass imaging plate.
3. Under a stereo microscope, use a pipette to remove as much medium as possible, being careful not to touch or damage the embryos; then, replace the medium with the previously prepared 1% low melting agarose solution. Next, use a plastic thin-tapered tip fixed to the end of a teasing needle to gently push each embryo to the bottom of the imaging plate, orienting it so that its dorsal side is facing the glass to ensure a good view of the optic tectum and a minimal amount of low melting agarose in between the embryo and the glass.
4. Image the mounted embryo promptly using an inverted confocal microscope featuring a high-quality Plan Apo Objective (10x/0.45 Plan Apo or a 20x/0.75 Plan Apo used here). If the timelapse will be running for an extended period (over an hour), add one or two drops of E3 buffer with Tricaine methanesulfonate (160 mg/L) to the bottom glass imaging plate to keep the low-melting agarose from drying excessively.
   NOTE: It is crucial to wait for the agarose to completely solidify before adding E3 buffer and tricaine methanesulfonate media to the imaging disk; otherwise, the fish could move slowly laterally and axially, thus causing a sample drift in the timelapse.
5. Proceed to set up the timelapse acquisition for the desired time: set the image resolution to 1,024 x 1,024, with a pixel size of 0.49 µm. Collect optical slices in 2-3 µm increments to ensure an acceptable resolution on the z-axis. In the zebrafish embryo, enclose the optic tectum in a 100-150 µm z-stack, depending on the developmental stage used. To ensure successful cell tracking, keep the time interval between frames between 30 s and 60 s.

3. Tracking analysis and data export

1. Convert the timelapse file into IMS format using the Imaris file converter software (see Table of Materials). Upon launching the software, either drag and drop the files into the input area or utilize the Add Files… button to select them manually. In the Output menu, designate the location for the converted files, opting for either the same folder as the input files or specifying a particular folder. Ensure the dimensions of the voxel size displayed are correct by clicking on the Set Voxel Size button and once the voxel size is verified, press the Start All button to start the file conversion (See Supplemental File 1 for a step-by-step guide of all the data analysis).
   NOTE: If the timelapse was performed on two or more embryos in parallel, the file will be automatically split into single positions, separating each embryo into different files. It is possible to use the software in batch mode and upload and convert multiple files at the same time. IMS files will be created and saved in the selected destination folder or the folder where the input files are located.
2. Once launched, let the analysis software start in the Arena; click on Observe Folder to open the previously converted file. Once the file is loaded into the software, use the Slice view to scroll through the z-stack, using the slider in the Slice Toolbar situated on the left of the View Area, and measure some cell diameters by drawing the diameter of the selected cells using the pointer. Click and drag the mouse to draw a segment spanning the cell core; observe the length of the segment drawn in the Measure Toolbar, situated on the right of the View Area.
   NOTE: Using the imaging settings in step 2.5, the cell body diameter can be expected to be between 6 µm and 8 µm.
3. To automatically detect the cells, go back to the 3D view and click on the spot icon in the Object Toolbar to add a new Spots object in the object list and open the automatic Spots creation wizard. In the first step of the wizard, tick the option to segment only a region of interest. Then, enable the Track Spots (over time) option to compute the tracking data, as well as the Object-Object Statistics to allow comparison between spots and expand the range of data that will be available later in the analysis. Finally, proceed in the wizard by clicking the Next button in the lower right of the wizard window.
4. In the second step, proceed to specify a region of interest (ROI) area that encloses the optic tectum of the embryo. Modify the size of the ROI by clicking and dragging the small white arrows present on each face of it. Once the ROI size is defined, extend it to all the recorded frames by adjusting the time interval.
5. Then, select as Source Channel the channel used to detect the cells, and set the cell diameter measurement, previously obtained in step 3.2, as the Estimated XY Diameter. Enable the Background Subtraction option and click on Next.
6. At this point in the Spot creation wizard, the software incorporates a quality filtering step wherein spots are filtered by the signal intensity measured at the center of each spot; in the channel where the spots were detected. Adjust the intensity threshold by entering the value directly in the data field for lower and upper threshold boxes or by dragging the corresponding-colored lines in the histogram shown at the bottom of the creation wizard, so that all cells are detected, including those localized deeper in the tissue, which usually show a lower signal intensity. All these changes are simultaneously visible in the View Area.
   NOTE: It is possible to observe multiple spots labeling a single cell at this point, but this can be corrected later in the analysis.
   In longer timelapses, if some photobleaching is seen, check the View Area of multiple frames before setting the threshold to make sure that all cells are also detected in the most recent and therefore, fainter frames.
7. Once satisfied, click on Next to validate the selected quality thresholding. All the spots satisfying the quality thresholding parameter and having a diameter matching with the Estimated XY Diameter will be labeled as a spot. Visualize this as a preview in the View Area, where spots satisfying these two parameters will be superimposed on the Source Channel.
8. In the next step, select the Autoregressive Motion as the tracking algorithm to track cells that have a more or less continuous motion. Judge the longest distance a cell moves between two time points by carefully observing the motion of the spots in between two frames and enter an estimated number in the Max Distance field. This is the distance that a spot is allowed to deviate from the predicted future position.
9. Set the Maximum Gap size to the least number of time points in which the spot may disappear without interrupting the track. If the interval between frames is below 60 s, use a size of 3. If the time interval between frames is longer, increase the maximum gap size allowed to reduce track fragmentation.
10. Enable the Fill gaps with all detected objects to drop the detection threshold close to the expected position predicted by the tracking algorithm and connect tracks that are interrupted by a failure of the spot detection. Finally, click on Next and let the software automatically generate tracks for all the previously generated spots.
11. If the obtained tracks appear to be too numerous or fragmented go back in the creation wizard, by clicking on the Previous button, and adjust the maximum distance previously defined, either increasing or decreasing the number until most of the obtained tracks are representative of the cell movements.
    NOTE: The number of tracks should roughly match the number of microglia cells, therefore, ranging between 30 and 50, depending on the embryo's age. However, before completion of the analysis, a higher number is expected due to erroneous tracks.
12. Once the tracks have been calculated, the software incorporates a filtering step to filter out all the very brief tracks as they are often inaccurate or not informative. Exclude tracks shorter than 3-8 minutes (min) by entering the value directly in the data field for the lower limit of the track duration filter, or by dragging the corresponding-colored lines in the histogram shown at the bottom of the creation wizard. Click on Next to validate the filter and move on in the creation wizard.
    NOTE: As mentioned before, by modifying the maximum (max) and minimum (min) track duration threshold, it is possible to preview the tracks that will be filtered and excluded from the analysis. The filtered tracks can then be duplicated in a new object for further analysis. The exact time threshold for filtering may vary based on the results of the previous creation wizard step. We recommend fine-tuning these filter settings by reviewing the preview in the View Area for optimal results.
13. Since the Tg(mpeg1:eGFP) transgenic line labels all mononuclear phagocytes, skin macrophages in addition to microglia will also be detected in timelapse movies. Manually remove the tracks generated from skin macrophages from the analysis to focus on brain parenchymal microglia (Figure 3).
14. Manually correct other potential mistakes in the tracks in the track editor, where each data point is displayed as an object in the track. Activate the circle select mode button on the right of the View Area and highlight tracks that require modifications or adjustments. Once the problematic track is selected, using the connect and disconnect buttons, edit it to correctly represent the cell movements. Use the track editor to also select and delete duplicated single spots or small track fragments that erroneously represent the same cell by selecting them with the circle select mode and pressing Canc or Delete to eliminate them.
    NOTE: If the settings of the tracking are correct, this step should be required for only a small percentage of the tracked cells.
15. If the microglia reporter line is used in combination with fluorescent transgenics labeling other cell types in the parenchyma, perform segmentation as a new surface object or spot object to visualize them. To do so, repeat the steps above (from steps 3.2 to 3.14), modifying the Source Channel input in step 3.5, for it to be the one used to image the second cell population, and taking care of adapting the descriptive parameters specific to the cells of interest such as Estimated XY Diameter (step 3.5), Max Distance (step 3.8) to the new cell population. Other steps can be repeated without changes.
    NOTE: In case of a new surface creation, Estimated XY Diameter is replaced by a thresholding option based on Background Subtraction, where the input to be given is the diameter of the largest sphere which fits into the object. This value can be estimated from the Slice view using the Measure toolbar as explained in step 3.2.
16. Once satisfied with the obtained tracks, extract the desired tracking statistics from the Statistic tab. Clicking on the settings button, select which statistics will be computed for each spot object or surface object previously created. In case more objects have been created in the same analysis, two spot objects or a spot and a surface object, statistics depicting the relative position of one object to the other, such as the shortest distance between the two, will also be available to export.
    NOTE: All the desired values can be either specific or average values, and all raw data are downloadable as a Comma Separated Values (CSV) file using the buttons at the bottom of the Statistik tab. Provided that the analysis also includes other cell types, additional statistics on the relative distance of one object to the other will also become available.