My research project focuses on characterizing microglial dynamics, morphology and function during the first week of zebrafish embryonic development. Microglia are highly dynamic cells and we believe that a deeper characterization of their behavioral patterns and morphological changes could give us an interesting insight into their maturation process and activity in the developing brain. The major challenge in this type of imaging lies in capturing multiple channels with high quality and an excellent temporary resolution.
However, advancement in confocal technology are now enabling us to meet this challenge effectively. We believe our protocol to be a potent tool that could facilitate the characterization of microglial dynamics in diverse settings. This includes not only different stages of development, but also very smooth under background where microglia homeostasis is compromised.
To begin, obtain anesthetized zebrafish embryos in E3 buffer. After selecting the embryos with the desired fluorescence, visualize their heartbeat under a bright field stereo microscope. Using a wide tip transfer pipette, draw up to three embryos and transfer them to a bottom glass imaging plate.
Under a stereo microscope, using a pipette, replace the medium around the embryos with a 1%low melting agao solution. Using a plastic thin tapered tip attached to a teasing needle, gently push each embryo to the bottom of the imaging plate and orient the dorsal side to face the glass. Mount the embryo in an inverted confocal microscope.
After setting the time lapse acquisition parameters, acquire the images. Open the Imaris file converter software, and drag and drop the files into the input area. In the output menu, designate the location for the converted files.
Click on the Set Voxel Size to verify and press the Start All button to start the file conversion to IMS format. 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, click the slice view to scroll through the c stack. Using the slider in the slice toolbar, measure cell diameters after drawing the diameter of the selected cells with the pointer. Click and drag the mouse to draw a segment spanning the cell core and observe the length of the segment drawn in the measure toolbar situated on the right of the view area.
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 overtime option to compute the tracking data as well as the objective object statistics to allow comparison between spots and expand the range of data.
Click the Next button in the lower right of the wizard window. Specify a region of interest or ROI area that encloses the optic tectum of the embryo. Click and drag the small white arrows present on each face of it to modify the size of the ROI.
Then extend it to all the recorded frames with time interval adjustments. Then select a source channel and set the cell diameter measurement obtained previously as the estimated XY diameter. Enable the background subtraction option and click on Next.
Enter the intensity threshold values directly into the lower and upper threshold boxes to ensure the detection of all cells. Click on View Area to observe all these changes in real time. Once satisfied, click on Next to validate the selected quality thresholding and click on View Area to visualize spots satisfying these two parameters superimposed on the source channel.
Next, select the auto aggressive motion as the tracking algorithm to track cells that have a more or less continuous motion. Observe the motion of the spots in between two frames to judge the longest distance a cell moves between two time points, and enter an estimated number in the max distance field. To set the maximum gap size, if the interval between frames is below 60 seconds, use a size of three, and for longer time intervals, increase the same.
Enable the fill gaps with all detected objects to drop the detection threshold and connect tracks. Then click on Next and let the software automatically generate tracks for all the previously generated spots. If the obtained tracks are numerous or fragmented, return to the creation wizard with the Previous button and adjust the maximum distance previously defined.
To exclude the tracks shorter than three to eight minutes, enter the value directly in the data field for the lower limit of the track duration filter. Click on Next to validate the filter and move on to creation wizard. Manually remove the tracks generated from skin macrophages from the analysis to focus on brain parenchymal microglia.
Using the track editor, manually correct other potential mistakes in the tracks. On the right of the view area, activate the circle select mode button 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.
With the track editor, select Circle Select Mode to delete duplicated single spots. If using microglia reporter with other fluorescently labeled parenchymal cells, adjust the estimated XY diameter and max distance to the new cell population. Modify the source channel input to the image.
Finally, using the statistic tab, extract the desired tracking statistics. Click on the settings button and select the statistics for the computation of each spot object or surface object previously created. The brain of the embryonic zebrafish was imaged in its entirety, and the location of neurons with respect to microglia was visualized.
The spatial data obtained using this protocol depicts the average speed of microglial cells, the average distance they cover in one hour, their mean square displacement and their distribution in space at different times.
