The authors would like to express their sincere gratitude to Professor Nicolas Bayens for generously providing access to the confocal microscope essential for this study.&#160;This work was funded in part by the Funds for Scientific Research (FNRS) under Grant Numbers F451218F and UG03019F, the Alzheimer Research Foundation (SAO-FRA) (to V.W.), A.M. is supported by a Research Fellowship from the FNRS. Figure 1&#160;was created on biorender.com.

Zebrafish were maintained under standard conditions, according to FELASA42 and institutional (Universit&#233; 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.
<ol>
	<li>Maintain zebrafish (Danio rerio) at a temperature of 28 &#176;C and manage them in accordance with established protocols11.</li>
	<li>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&#39; birth date. Remove the separator the following morning to time the mating of the fish.</li>
	<li>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 Casperw2, that allows for clear imaging without the need to expose embryos to PTU for an extended period of time13.</li>
	<li>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).
	<ol>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>
2. Zebrafish mounting
<ol>
	<li>Prepare a 1% low melting agarose solution in E3 buffer. Cool the agarose to 37 &#176;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.</li>
	<li>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&#39;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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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 &#181;m. Collect optical slices in 2-3 &#181;m increments to ensure an acceptable resolution on the z-axis. In the zebrafish embryo, enclose the optic tectum in a 100-150 &#181;m 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.</li>
</ol>
3. Tracking analysis and data export
<ol>
	<li>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&#8230;&#160;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.</li>
	<li>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 &#181;m and 8 &#181;m.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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&#39;s age. However, before completion of the analysis, a higher number is expected due to erroneous tracks.</li>
	<li>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.</li>
	<li>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).</li>
	<li>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.</li>
	<li>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.</li>
	<li>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 Statistics 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.</li>
</ol>

Elucidating Microglia Behavior in Zebrafish Optic Tectum Development

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(157), e60593 (2020).</li><li>Tayanloo-Beik, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+modeling+of+autism+spectrum+disorders,+current+status+and+future+prospective.">Zebrafish modeling of autism spectrum disorders, current status and future prospective.</a> Front Psychiatry. 13, 911770 (2022).</li><li>D'Amora, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+as+an+innovative+tool+for+epilepsy+modeling:+State+of+the+art+and+potential+future+directions.">Zebrafish as an innovative tool for epilepsy modeling: State of the art and potential future directions.</a> Int J Mol Sci. 24 (9), 7702 (2023).</li><li>Langova, V., Vales, K., Horacek, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+zebrafish+and+laboratory+rodents+in+schizophrenia+research.">The role of zebrafish and laboratory rodents in schizophrenia research.</a> Front Psychiatry. 11, 549232 (2020).</li><li>Chia, K., Klingseisen, A., Sieger, D., Priller, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+as+a+model+organism+for+neurodegenerative+disease.">Zebrafish as a model organism for neurodegenerative disease.</a> Front Mol Neurosci. 15, 940484 (2022).</li><li>Astell, K. R., Sieger, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+in+vivo+models+of+cancer+and+metastasis.">Zebrafish in vivo models of cancer and metastasis.</a> Cold Spring Harb Perspect Med. 10 (8), 037077 (2020).</li></ol>

The authors declare no conflicts of interest.

The current protocol enables in vivo imaging of microglia dynamics in a vertebrate embryo and visualization of the acquired motility data. Microglia colonization of the developing brain occurs very early during embryogenesis and precedes critical events such as peaks of neurogenesis, astrogliogenesis, oligodendrogenesis, and many other cellular processes17. It is therefore not surprising that microglia play important functions in shaping specific aspects of brain development18, for example, through the regulation of neuronal differentiation, migration, and survival19,20,21, as well as synaptic pruning5&#160;and myelination22,23,24.
The contribution of dysfunctional microglia to the pathogenesis and/or progression of neurodevelopmental disorders is also increasingly being recognized25. Indeed, the early presence of microglia in the forming brain exposes these cells to distinct physiological states26&#160;and environmental changes. This can have a significant impact given that microglia are long-lived cells in both rodents and humans, being maintained during the lifespan through self-renewal of local progenitors27,28,29. We believe this protocol could serve as a powerful tool to better characterize the behavior of microglia in these distinct physiological states, as they develop, mature, and establish their network during the successive steps of brain morphogenesis.
Using the setup described here, we have successfully imaged and acquired data on zebrafish larvae as old as 6 dpf. Extending the analysis to later developmental stages will likely succeed but will require adjusting the imaging setup to take into consideration the increased sample size, especially along the z-axis. In attempting this, we suggest focusing on maintaining a low signal-to-noise ratio and fast scanning time, as they are key parameters for a successful analysis.
We suggest a minimum imaging time of 1 h to allow for microglia tracking; the longest imaging window that has been tested with this protocol is 8 h. Moreover, it is important for the tracking analysis to keep the time interval between frames as short as possible, ideally between 30 s and 60 s. This will allow for more accurate and detailed tracking data in downstream analyses. Therefore, especially if detecting more than one fluorophore, it is fundamental to avoid spectral overlap and ensure enough separation between the two fluorophore emission spectra to allow for a simultaneous acquisition, without signal bleedthrough.
Other protocols for high-quality timelapse recording of the zebrafish brain are available30, but this is the first one showing how to successfully track all microglia movement during embryonic development over an extended period. Although the workflow presented here focused on tracking microglia in a physiological context, it can be easily applied to the analysis of microglia in pathology. Indeed, several models of neurodevelopmental disorders, such as autism31, epilepsy32, and schizophrenia33, but also neurodegeneration34&#160;and cancer35, have been established in zebrafish that provide unique opportunities for determining microglial response and behavior in disease conditions.
Notably, this tracking protocol is highly versatile and could also be instrumental for shedding light on the migration patterns of various cell types across diverse anatomical regions of the zebrafish embryo, thus potentially opening up avenues for additional applications, beyond the microglial investigation scope described in this article. Moreover, by harnessing the ability to combine multiple fluorescent transgenic lines, we gain the ability to discern the spatial relationship between microglia and other cell types of the brain microenvironment, with the potential to visualize cellular interactions and cross-talks throughout timelapse recordings, in a non-invasive way. This could be instrumental in unraveling the physiological significance of microglia behavior and contribute to a deeper characterization of these highly motile cells.

As resident macrophages in the central nervous system (CNS), microglia represent a distinct non-neuronal population that accounts for up to 15% of all glial cells in the adult brain. Studying microglia biology has gained increasing attention in recent years due to their established importance in development, physiology, and disease1. Under physiological conditions, microglial cells are highly dynamic, continuously surveying the brain parenchyma2,3. This behavior allows microglia to colonize the brain and play pivotal roles in its development such as shaping neuronal circuitry4, synaptic pruning5, and vasculogenesis6. Furthermore, this inherent dynamic nature allows microglia to constantly monitor the CNS for signs of infection, injury, or any deviations from homeostasis7. To dissect these intricate cell dynamics, live imaging of microglia across space and time is indispensable. Fortunately, the optical transparency of externally developing zebrafish embryos, coupled with the availability of well-characterized transgenic reporter lines that fluorescently label microglia, positions zebrafish as an ideal vertebrate model for such investigations. Live imaging in zebrafish embryos offers a non-invasive approach that does not require surgery or extensive tissue manipulation, minimizing potential perturbations to the CNS status. This is a critical consideration when studying microglial cells, as they are highly sensitive to even subtle changes in the extracellular environment8.
Here, we provide a guideline to successfully track 3D microglial cell movements in the zebrafish embryo, allowing for an unprecedented view of microglia behavior within the intact architecture of the developing brain parenchyma (see Figure 1&#160;for a graphical overview of the protocol). This step-by-step protocol details how to set up and image zebrafish microglia at different developmental stages and how to extract high-resolution data on microglia cell motility to provide valuable insights into their migratory patterns and responses to environmental cues. We also demonstrate this protocol can be adapted to perform live multi-color imaging, thus extending its applicability to studying microglia in combination with transgenic lines that mark neighboring cells, including neurons3, oligodendrocytes9, and endothelial cells10&#160;(as shown in Figure 2). By adding to the toolbox that allows one to directly observe and characterize the dynamics of microglia behavior&#160;in real time and in their natural environment, this protocol will likely contribute to better elucidating microglia functionality during early development, both in physiology and disease.

<table><tbody><tr><td>1 L Breeding tanks</td><td>Tecniplast</td><td>ZB10BTE</td><td></td></tr><tr><td>1-phenyl-2-thiourea (PTU)</td><td>Sigma-Aldrich</td><td>P7629</td><td>Diluted to 0.2 mM in E3 to prevent embryo pigmentation</td></tr><tr><td>Bottom glass imaging dish</td><td>FluoroDish</td><td>FD3510-100</td><td></td></tr><tr><td>Disposable Graduated transfer pipette</td><td>avantor</td><td>16001-188</td><td></td></tr><tr><td>Dry block heater</td><td>Novolab</td><td>Grant QBD4</td><td>To keep low melting agarose at 37 &amp;deg;C</td></tr><tr><td>Ethyl 3-aminobenzoate methanesulfonate (Tricaine)</td><td>Sigma-Aldrich</td><td>E10521-50G</td><td></td></tr><tr><td>Imaris 10.0</td><td>Oxford Instruments</td><td></td><td>analysis software</td></tr><tr><td>Imaris File Converter</td><td>Oxford Instruments</td><td></td><td>https://imaris.oxinst.com/big-data</td></tr><tr><td>Laser-scanning confocal microscope</td><td>Nikon</td><td>Eclipse Ti2-E</td><td></td></tr><tr><td>Methylene blue</td><td>Sigma-Aldrich</td><td>M9140-25G</td><td></td></tr><tr><td>microloader tips</td><td>Eppendorf</td><td>5242956003</td><td></td></tr><tr><td>NuSieve GTG Agarose</td><td>Lonza</td><td>&amp;nbsp;50081</td><td></td></tr><tr><td>Petri dishes (90 mm)</td><td>avantor</td><td>391-0559</td><td></td></tr><tr><td>Pronase</td><td>Sigma-Aldrich</td><td>11459643001</td><td></td></tr><tr><td>&amp;nbsp;Stainless Steel Forceps Dumont No. 5</td><td>FineScienceTools</td><td>11254-20</td><td></td></tr><tr><td>Stereo microscope</td><td>Leica</td><td>Leica M80</td><td>To mount the embryos</td></tr><tr><td>teasing needle</td><td>avantor</td><td>76549-024</td><td></td></tr></tbody></table>

live imaging and characterization of microglia dynamics in the zebrafish embryo

Microglia are highly dynamic cells and their migration and colonization of the brain parenchyma is a crucial step for proper brain development and function. Externally developing zebrafish embryos possess optical transparency, which along with well-characterized transgenic reporter lines that fluorescently label microglia, make zebrafish an ideal vertebrate model for such studies. In this paper, we take advantage of the unique features of the zebrafish model to visualize the dynamics of microglia cells in vivo and under physiological conditions. We use confocal microscopy to record a timelapse of microglia cells in the optic tectum of the zebrafish embryo and then, extract tracking data using the IMARIS 10.0 software to obtain the cells' migration path, mean speed, and distribution in the optic tectum at different developmental stages. This protocol can be a useful tool to elucidate the physiological significance of microglia behavior in various contexts, contributing to a deeper characterization of these highly motile cells.

Microglial cells expressing Green fluorescent protein (eGFP) and endothelial cells expressing DsRed in Tg(mpeg1:eGFPgl22; kdrl:cres898; actb2:loxP-STOP-loxP-DsRedexpress,sd5) triple transgenic embryos14 were imaged at 3 dpf, according to the described protocol. A single zebrafish embryo was mounted in 1% low melting agarose on a bottom glass plate and the imaging process did not hinder the embryo&#39;s growth during the acquisition time. The timelapse was recorded using a commercial point-scanning confocal microscopy system equipped with a 10x 0.45 NA dry objective lens, and 488 nm and 561 nm excitation lasers were used for imaging microglia and endothelial cells, respectively. In addition, the time interval, image resolution, pixel size, and z step were 30 seconds (s), 1024x1024, 0.49 &#181;m, and 2.5 &#181;m, respectively. The timelapse recording lasted 3.5 hours (h).
The acquired field of view covered the whole head of the embryo, but the analysis focused specifically on the optic tectum, as during the first week of development, microglia are mainly restricted to the neuronal soma layer of this region of the dorsal midbrain, allowing investigators to visualize the whole population simultaneously15. The 3D tracking analysis was performed using Imaris 10.0, as explained above. As shown in Figure 4, the tracking was successful, resulting in 25 tracks, matching the expected number of microglia cells present in the optic tectum at 3 dpf16. A minimal manual tracks correction was required. Figure 5 shows an example of the data that is possible to extract from a successful tracking experiment. The simultaneous labeling of macrophages and endothelial cells allows for the quantification of microglia&#39;s position relative to the latter, allowing researchers to visualize each cell&#39;s relative distance to the closest endothelial cell in time and examine the frequency and number of potential interactions (Figure 5).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66533/66533fig01.jpg" /> 
Figure 1: Overview of the experimental procedure. (A) Zebrafish embryo preparation and anesthesia. (B) Sample mounting and positioning. (C) Image acquisition. (D) Image processing and extraction of motility data. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66533/66533fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66533/66533fig02.jpg" /> 
Figure 2: Microglia imaging and interactions with the brain cellular environment in the zebrafish embryo. (A) Brightfield image of a 3 dpf embryonic zebrafish head and brain (dorsal view), labeling the optic tectum and hindbrain. The brain of the embryonic zebrafish can be imaged in its entirety at this stage due to its small size and optical transparency. (B-D) Microglia location and behavior can be visualized by the use of transgenic lines such as (B,C) Tg(mpeg1:eGFP)gl22 and (D) Tg(mpeg1:mcherry)gl23, and focusing on the OT. (B) Neurons and their cell bodies can be identified using the reporter line Tg(XlTubb:DsRed)zf148 and microglia-neuron interactions can be visualized by merging the two channels in lines that co-express both transgenes. Seen is a merge of the green (microglia) and red (neurons, in magenta) signals. (C,D) Reporter lines can also shed light on microglia (in green) interactions with endothelial cells, here labeled in red using the (C) double Tg(kdrl:cres898; actb2:LOXP-STOP-LOXP-Dsredexpress, sd5) transgenics, or with glial cells such as oligodendrocytes and their precursors, using the (D) double Tg(olig2:EGFP; mpeg1:mcherry ) transgenic line. 1: oligodendrocytes and progenitor cells in the OT, 2: eurydendroid neurons in the cerebellum. Scale bars = 50 &#181;m. Abbreviations: dpf = days post fertilization; OT = optic tectum; HB = hindbrain. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66533/66533fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66533/66533fig03.jpg" /> 
Figure 3: Morphological and spatial disparity between skin macrophages and microglia in a 3 dpf zebrafish embryo. (A,B) Dorsal view of a Tg(mpeg1:eGFP)gl22 zebrafish embryo at 3 dpf, depicting (A) parenchymal microglia cells (marked by magenta asterisks) versus skin macrophages (white asterisks), identified based on their relative position along the z-axis, as seen in (B), which shows the monochrome 3D image rendered with z-axis color-coding. (C,D) Lateral view of (C) A and (D) B, showcasing the z-depths of mpeg1:eGFP+ cells within the embryo&#39;s head and highlighting the superficial localization of skin macrophages as compared to microglial cells. (E-K) High magnification of each cell indicated by an asterisk in A and B, providing a detailed visualization of the distinct morphologies between (E-G) amoeboid microglia and (H-K) elongated skin macrophages. Scale bars= 30 &#181;m. Abbreviation: dpf = days post fertilization. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66533/66533fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66533/66533fig04.jpg" /> 
Figure 4: Tracking of microglial cells in a 3 dpf zebrafish embryo. (A) Head dorsal view of a 3 dpf Tg(kdrl:cres898; actb2:LOXP-STOP-LOXP-Dsredexpress,sd5;mpeg1:eGFPgl22) triple transgenic embryo, showing microglia (green) and the surface rendering of the vessels (magenta). (B-D) Representative tracking of microglia movements over a (B) 3.5 h time window. Individual cells are seen to follow complex trajectories. (C) Detail from timelapse, showing a magnified view of the region in B enclosed by the dashed square. Six frames from the movie (45 min apart) are presented, documenting microglia establishing transient contacts with endothelial cells in their microenvironment. (D) Trajectories of individual microglial cell migration within the optic tectum, with the X, Y, and Z axes representing spatial dimensions. Scale bar = 50 &#181;m. Abbreviation: dpf = days post fertilization. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66533/66533fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66533/66533fig05.jpg" /> 
Figure 5: Visualization of the data obtained. Example of the spatial data that can be obtained using the described protocol. The graphs depict (A) the average speed of microglial cells, (B) the average distance they cover in 1 h, (C) their mean square displacement, and (D) their distribution in space at different times. The vessel&#39;s surface rendering also allowed for the measurement of the shortest distance between microglia and endothelial cells at any given time, both at (E) single-cell level and as a (F) global average. For A,B, and D, each dot represents an individual cell. N = one embryo. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66533/66533fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental File 1: <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66533/66533_20240306_SupplementalFile1 (2).zip" target="_blank">Please click here to download this File.</a>

Read this Scientific Article Protocol about Live Imaging and Characterization of Microglia Dynamics in the Zebrafish Embryo at JoVE.com

Author Spotlight: Using Zebrafish to Explore Microglia Migration During Brain Development

We demonstrate a method that takes advantage of fast scanning confocal microscopy to perform live imaging of microglia cells in the developing zebrafish optic tectum, allowing for the analysis of the dynamics of these cells in vivo.

Live Imaging and Characterization of Microglia Dynamics in the Zebrafish Embryo

Microglia are highly dynamic cells and their migration and colonization of the brain parenchyma is a crucial step for proper brain development and ...

live-imaging-characterization-microglia-dynamics-zebrafish

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

1.1K Views.  Université Libre de Bruxelles (ULB). Mijn onderzoeksproject richt zich op het karakteriseren van de dynamiek, morfologie en functie van microglia tijdens de eerste week van de embryonale ontwikkeling van zebravissen. Microglia zijn zeer dynamische cellen en we geloven dat een diepere karakterisering van hun gedragspatronen en morfologische veranderingen ons een interessant inzicht zou kunnen geven in hun rijpingsproces en activiteit in de zich ontwikkelende hersenen. De grootste uitdaging bij di...

Video: Auteur in de schijnwerpers: zebravissen gebruiken om microglia-migratie tijdens de ontwikkeling van de hersenen te onderzoeken

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Developmental Biology

Culture of Adult Transgenic Zebrafish Retinal Explants for Live-cell Imaging by Multiphoton Microscopy

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The M&#252;ller glia re-enter the cell cycle and produce neuronal progenitor cells that undergo subsequent rounds of cell divisions and differentiate into the lost neuronal cell types. Both M&#252;ller glia and neuronal progenitor cell nuclei replicate their DNA and undergo mitosis in distinct locations of the retina, i.e. they migrate between the basal Inner Nuclear Layer (INL) and the Outer Nuclear Layer (ONL), respectively, in a process described as Interkinetic Nuclear Migration (INM). INM has predominantly been studied in the developing retina. To examine the dynamics of INM in the adult regenerating zebrafish retina in detail, live-cell imaging of fluorescently-labeled M&#252;ller glia/neuronal progenitor cells is required. Here, we provide the conditions to isolate and culture dorsal retinas from Tg[gfap:nGFP]mi2004 zebrafish that were exposed to constant intense light for 35 h. We also show that these retinal cultures are viable to perform live-cell imaging experiments, continuously acquiring z-stack images throughout the thickness of the retinal explant for up to 8 h using multiphoton microscopy to monitor the migratory behavior of gfap:nGFP-positive cells. In addition, we describe the details to perform post-imaging analysis to determine the velocity of apical and basal INM. To summarize, we established conditions to study the dynamics of INM in an adult model of neuronal regeneration. This will advance our understanding of this crucial cellular process and allow us to determine the mechanisms that control INM.

Zebrafish retinal regeneration has mostly been studied using fixed retinas. However, dynamic processes such as interkinetic nuclear migration occur during the regenerative response and require live-cell imaging to investigate the underlying mechanisms. Here, we describe culture and imaging conditions to monitor Interkinetic Nuclear Migration (INM) in real-time using multiphoton microscopy.

An endogenous regeneration program is initiated by M&#252;ller glia in the adult zebrafish (Danio rerio) retina following neuronal damage and death. The ...

Multi-Photon Time Lapse Imaging to Visualize Development in Real-time: Visualization of Migrating Neural Crest Cells in Zebrafish Embryos

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to numerous cell types throughout the body. Understanding the biology of the neural crest has been limited, reflecting a lack of genetically tractable models that can be studied in vivo and in real-time. Zebrafish is a particularly important developmental model for studying migratory cell populations, such as the neural crest. To examine neural crest migration into the developing eye, a combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time videos of the developing eye in transgenic zebrafish embryos, namely Tg(sox10:EGFP) and Tg(foxd3:GFP), as sox10 and foxd3 have been shown in numerous animal models to regulate early neural crest differentiation and likely represent markers for neural crest cells. Multi-photon time-lapse imaging was used to discern the behavior and migratory patterns of two neural crest cell populations contributing to early eye development. This protocol provides information for generating time-lapse videos during zebrafish neural crest migration, as an example, and can be further applied to visualize the early development of many structures in the zebrafish and other model organisms.

A combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time imaging of neural crest migration in Tg(sox10:EGFP) and Tg(foxd3:GFP) zebrafish embryos.

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to ...

Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning fundamental developmental biology through to large-scale genetic and chemical screens. However, many experiments, especially those related to infection and xenograft models, rely on microinjection and imaging of embryos and larvae, which are laborious techniques that require skill and expertise. To improve the precision and throughput of current microinjection techniques, we developed a series of microstructured devices to orient and stabilize zebrafish embryos at 2 days post fertilization (dpf) in ventral, dorsal, or lateral orientation prior to the procedure. To aid in the imaging of embryos, we also designed a simple device with channels that orient 4 zebrafish laterally in parallel against a glass cover slip. Together, the tools that we present here demonstrate the effectiveness of photolithographic approaches to generate useful devices for the optimization of zebrafish techniques.

Microinjection of zebrafish embryos and larvae is a crucial but challenging technique used in many zebrafish models. Here, we present a range of microscale tools to aid in the stabilization and orientation of zebrafish for both microinjection and imaging.

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Biology

Live Imaging of Cell Motility and Actin Cytoskeleton of Individual Neurons and Neural Crest Cells in Zebrafish Embryos

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily accessible at all stages of development. Moreover, their optical clarity allows high resolution imaging of cell and molecular dynamics in the natural environment of the intact embryo. We are using a live imaging approach to analyze cell behaviors during neural crest cell migration and the outgrowth and guidance of neuronal axons.
Live imaging is particularly useful for understanding mechanisms that regulate cell motility processes. To visualize details of cell motility, such as protrusive activity and molecular dynamics, it is advantageous to label individual cells. In zebrafish, plasmid DNA injection yields a transient mosaic expression pattern and offers distinct benefits over other cell labeling methods. For example, transgenic lines often label entire cell populations and thus may obscure visualization of the fine protrusions (or changes in molecular distribution) in a single cell. In addition, injection of DNA at the one-cell stage is less invasive and more precise than dye injections at later stages.
Here we describe a method for labeling individual developing neurons or neural crest cells and imaging their behavior in vivo. We inject plasmid DNA into 1-cell stage embryos, which results in mosaic transgene expression. The vectors contain cell-specific promoters that drive expression of a gene of interest in a subset of sensory neurons or neural crest cells. We provide examples of cells labeled with membrane targeted GFP or with a biosensor probe that allows visualization of F-actin in living cells1.
Erica Andersen, Namrata Asuri, and Matthew Clay contributed equally to this work.

This protocol describes imaging of individual neurons or neural crest cells in living zebrafish embryos. This method is used to examine cellular behaviors and actin localization using fluorescence confocal time-lapse microscopy.

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily ...

Multicolor Time-lapse Imaging of Transgenic Zebrafish: Visualizing Retinal Stem Cells Activated by Targeted Neuronal Cell Ablation

High-resolution time-lapse imaging of living zebrafish larvae can be utilized to visualize how biological processes unfold (for review see 1). Compound transgenic fish which express different fluorescent reporters in neighboring cell types provide a means of following cellular interactions 2 and/or tissue-level responses to experimental manipulations over time. In this video, we demonstrate methods that can be used for imaging multiple transgenically labeled cell types serially in individual fish over time courses that can span from minutes to several days. The techniques described are applicable to any study seeking to correlate the "behavior" of neighboring cells types over time, including: 1) serial 'catch and release' methods for imaging a large number of fish over successive days, 2) simplified approaches for separating fluorophores with overlapping excitation/emission profiles (e.g., GFP and YFP), 3) use of hypopigmented mutant lines to extend the time window available for high-resolution imaging into late larval stages of development, 4) use of membrane targeted fluorescent reporters to reveal fine morphological detail of individual cells as well as cellular details in larger populations of cells, and 5) a previously described method for chemically-induced ablation of transgenically targeted cell types; i.e., nitroreductase (NTR) mediated conversion of prodrug substrates, such as metronidazole (MTZ), to cytotoxic derivatives 3,5.
As an example of these approaches, we will visualize the ablation and regeneration of a subtype of retinal bipolar neuron within individual fish over several days. Simultaneously we will monitor several other retinal cell types, including neighboring non-targeted bipolar cells and potential degeneration-stimulated retinal stem cells (i.e., M&#971;ller glia). This strategy is being applied in our lab to characterize cell- and tissue-level (e.g., stem cell niche) responses to the selective loss and regeneration of targeted neuronal cell types.

In this video, techniques for multicolor confocal time-lapse imaging and targeted cell ablation are provided. Time-lapse imaging is used to monitor the behavior of multiple cell types of interest in vivo. Targeted cell ablation facilitates the study neural circuit function and cell-specific neuronal regeneration paradigms.

High-resolution time-lapse imaging of living zebrafish larvae can be utilized to visualize how biological processes unfold (for review see 1). Compound ...

Long-term Live Imaging Device for Improved Experimental Manipulation of Zebrafish Larvae

The zebrafish larva is an important model organism for both developmental biology and wound healing. Further, the zebrafish larva is a valuable system for live high-resolution microscopic imaging of dynamic biological phenomena in space and time with cellular resolution. However, the traditional method of agarose encapsulation for live imaging can impede larval development and tissue regrowth. Therefore, this manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which was designed and fabricated as a functionally compartmentalized device to orient larvae for high-resolution microscopy while permitting caudal fin transection within the device and subsequent unrestrained tail development and re-growth. This device allows for wounding and long-term imaging while maintaining viability. Given that the zWEDGI mold is 3D printed, the customizability of its geometries make it easily modified for diverse zebrafish imaging applications. Furthermore, the zWEDGI offers numerous benefits, such as access to the larva during experimentation for wounding or for the application of reagents, paralleled orientation of multiple larvae for streamlined imaging, and reusability of the device.

This manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which is a compartmentalized device designed to orient and restrain zebrafish larvae. The design permits tail transection and long-term collection of high-resolution fluorescent microscopy images of wound healing and regeneration.

The zebrafish larva is an important model organism for both developmental biology and wound healing. Further, the zebrafish larva is a valuable system for ...

Bioengineering

In vivo Imaging of Fully Active Brain Tissue in Awake Zebrafish Larvae and Juveniles by Skull and Skin Removal

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at cellular and subcellular resolution. Advances in molecular and imaging technologies have allowed us to gain numerous detailed insights into cellular and molecular mechanisms of brain development in the transparent zebrafish embryo. Recently, processes of refinement of neuronal connectivity that occur at later larval stages several weeks after fertilization, which are for example control of social behavior, decision making or motivation-driven behavior, have moved into focus of research. At these stages, pigmentation of the zebrafish skin interferes with light penetration into brain tissue, and solutions for embryonic stages, e.g., pharmacological inhibition of pigmentation, are not feasible anymore.
Therefore, a minimally invasive surgical solution for microscopy access to the brain of awake zebrafish is provided that is derived from electrophysiological&#160;approaches. In teleosts, skin and soft skull cartilage can be carefully removed by micro-peeling these layers, exposing underlying neurons and axonal tracts without damage. This allows for recording neuronal morphology, including synaptic structures and their molecular contents, and the observation of physiological changes such as Ca2+ transients or intracellular transport events. In addition, interrogation of these processes by means of pharmacological inhibition or optogenetic manipulation is feasible. This brain exposure approach provides information about structural and physiological changes in neurons as well as the correlation and interdependence of these events in live brain tissue in the range of minutes or hours. The technique is suitable for in vivo brain imaging of zebrafish larvae up to 30 days post fertilization, the latest developmental stage tested so far. It, thus, provides access to such important questions as synaptic refinement and scaling, axonal and dendritic transport, synaptic targeting of cytoskeletal cargo or local activity-dependent expression. Therefore, a broad use for this mounting and imaging approach can be anticipated.

Here we present a method to image the zebrafish embryonic brain in vivo upto larval and juvenile stages. This microinvasive procedure, adapted from electrophysiological approaches, provides access to cellular and subcellular details of mature neuron and can be combined with optogenetics and neuropharmacological studies for characterizing brain function and drug intervention.

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at ...

Neuroscience

In Vivo Whole-Brain Imaging of Zebrafish Larvae

In Vivo Whole-Brain Imaging of Zebrafish Larvae Using Three-Dimensional Fluorescence Microscopy

As a vertebrate model animal, larval zebrafish are widely used in neuroscience and provide a unique opportunity to monitor whole-brain activity at the cellular resolution. Here, we provide an optimized protocol for performing whole-brain imaging of larval zebrafish using three-dimensional fluorescence microscopy, including sample preparation and immobilization, sample embedding, image acquisition, and visualization after imaging. The current protocol enables in vivo imaging of the structure and neuronal activity of a larval zebrafish brain at a cellular resolution for over 1 h using confocal microscopy and custom-designed fluorescence microscopy. The critical steps in the protocol are also discussed, including sample mounting and positioning, preventing bubble formation and dust in the agarose gel, and avoiding motion in images caused by incomplete solidification of the agarose gel and paralyzation of the fish. The protocol has been validated and confirmed in multiple settings. This protocol can be easily adapted for imaging other organs of a larval zebrafish.

Author Spotlight: In Vivo Whole-Brain Imaging of Zebrafish Larvae Using Three-Dimensional Fluorescence Microscopy  

Presented here is a protocol for in vivo whole-brain imaging of larval zebrafish using three-dimensional fluorescence microscopy. The experimental procedure includes sample preparation, image acquisition, and visualization.

As a vertebrate model animal, larval zebrafish are widely used in neuroscience and provide a unique opportunity to monitor whole-brain activity at the ...

Live Imaging and Characterization of Microglia Dynamics in the Zebrafish Embryo

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