The zebrafish lines used for calcium imaging were provided by the Zebrafish Center for Disease Modeling (ZCDM), Korea. This research was supported by National Research Foundation of Korea (2020R1C1C1009869, NRF2021R1A4A102159411, RS-2023-00209473).

All zebrafish experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of KAIST (KA2021-125). A total of 12 adult zebrafish with pan-neuronal expression of the calcium indicator GCaMP7a [Tg(huc:GAL4);Tg(UAS:GCaMP7a)] on a casper [mitfa(w2/w2);mpv17(a9/a9)] background were used for breeding. This group consisted of eight females and four males, with ages varying from 3 to 12 months. Imaging experiments were performed on larval zebrafish at 3-4 days post fertilization (d.p.f.), a stage during which their sex cannot be determined.
1. Zebrafish sample preparation
<ol>
	<li>Collect embryos after breeding adult fish of a desired transgenic line, such as Tg(huc:GAL4);Tg(UAS:GCaMP7a)20,21,22, in a Petri dish filled with egg water (see Table of Materials). Place the embryos in an incubator at 28 &#176;C and raise them to 3-4 d.p.f. larvae23,24,25.
	<ol>
		<li>If the zebrafish background is not albino, to inhibit a formation of pigmentation, transfer the embryos at 24 h post fertilization (h.p.f.) into the Petri dish filled with egg water containing 200 &#181;M of 1-phenyl 2-thiourea (PTU; see Table of Materials)25,26. Every 24 h, transfer the fish into a new dish with fresh egg water containing 200 &#181;M of PTU. 
		NOTE: The zebrafish are maintained under standard conditions at 28 &#176;C and a 14:10 h light:dark cycle. The PTU treatment is known to affect the behavior and thyroid function of larval zebrafish27,28. Therefore, it is important to use the PTU with caution and to carefully control for potential confounding factors in any experiments.</li>
	</ol>
	</li>
	<li>To find the sample expressing fluorescent proteins of interest (e.g., pan-neuronal GCaMP7a), screen the sample under an epi-fluorescence microscope and select a sample with a bright expression.</li>
	<li>Prepare the selected 3-4 d.p.f. zebrafish sample in the Petri dish filled with egg water (Figure 2A). 
	NOTE: To record spontaneous neuronal activity, it is recommended to use samples aged between 80-100 h.p.f., as the level of spontaneous activity is low before 80 h.p.f. and pigmentation development, even with a casper background, may degrade image quality after 100 h.p.f.</li>
	<li>Prepare a 0.25 mg/mL pancuronium bromide solution14,19,29,30,31 by adding 1 mL of a 2.5 mg/mL stock solution (see Table of Materials) to 10 mL of the egg water. Aliquot the pancuronium bromide solution into 1.5 mL microcentrifuge tubes.</li>
	<li>Transfer the pre-screened sample into the Petri dish using a transfer pipette. 
	NOTE: Try to carry over a minimum volume of egg water with the sample.</li>
	<li>Transfer 0.1 mL of the pancuronium bromide solution into the Petri dish for paralyzation. 
	&#8203;NOTE: Pancuronium bromide has a potential dampening effect on neural activity in larval zebrafish32. It is essential to consider the concentration and duration of exposure to pancuronium bromide carefully.</li>
</ol>
2. 2% (wt/vol) agarose gel preparation
<ol>
	<li>Turn on a heat block and set the target temperature to 37 &#176;C. Wait for the unit to heat up and equilibrate at the set temperature.</li>
	<li>Dissolve 0.2 g of low-melting point agarose powder (see Table of Materials) in 10 mL of egg water.</li>
	<li>Heat the agarose solution in a microwave and mix it by shaking and vortexing until the agarose is completely dissolved.</li>
	<li>Aliquot the agarose gel into 1.5 mL microcentrifuge tubes and store the microcentrifuge tubes on the heat block (Figure 2B). 
	&#8203;NOTE: Check if the bubbles in the agarose gel have disappeared.</li>
</ol>
3. Sample mounting and positioning
<ol>
	<li>Check the sample under a stereomicroscope to verify that larvae movement has stopped and to evaluate the sample&#39;s health visually by checking its heartbeat (Figure 2C). If the heartbeat is too slow, discard the sample. 
	NOTE: If the heartbeat of the fish being imaged is too slow (e.g., below 60 beats per minute), it may not be possible to perform long-term imaging. To assess the wellness of the fish, the heart rate can be visually checked by comparing it to other fish in the same Petri dish. This will help ensure that the fish being imaged is healthy and stable enough for the imaging procedure.</li>
	<li>Using the transfer pipette, place a single larval zebrafish into the agarose gel in the 1.5 mL microcentrifuge tube (Figure 2D). 
	NOTE: Ensure to discard the pipette after transferring the sample.</li>
	<li>Pour the agarose gel into the Petri dish to make a 1-2 mm coat. Transfer the sample in the microcentrifuge tubes to the Petri dish using the transfer pipette, so that the larva is placed in the center of the dish. 
	NOTE: If there is dust and bubbles in the agarose gel, remove them using the transfer pipette.</li>
	<li>Use forceps to position the sample in the desired orientation so that the head and tail are flat (Figure 2E).</li>
	<li>Rotate the sample using forceps so that both eyes are level (Figure 2F). 
	NOTE: Aligning procedures (position and rotation) should be completed before the agarose gel starts to solidify.</li>
	<li>After the alignment, wait until the agarose gel has solidified (Figure 2G,H). 
	NOTE: The waiting time for the agarose gel to solidify can range from 5-10 min, depending on the volume and size of the gel.</li>
	<li>After solidification of the agarose gel, fill the Petri dish with egg water and place the Petri dish with the embedded sample on the microscope stage (Figure 2I).</li>
</ol>
4. Image acquisition
<ol>
	<li>Turn on the microscope system (e.g., lasers, confocal controllers, microscope, and computer; see Table of Materials) and verify that the whole system is working.</li>
	<li>Select a low-magnification objective lens and locate the sample at the center of the field of view.</li>
	<li>Select a water-immersion or water-dipping objective lens with proper magnification (e.g., 16x 0.8 numerical aperture (NA) water-dipping lens; see Table of Materials). Make fine adjustments to the field of view.</li>
	<li>Set the imaging parameters (e.g., image size, laser power, exposure time, number of frames) using image acquisition software. 
	NOTE: Set the imaging parameters to achieve the best possible results for specific needs (see the setup for image acquisition in the representative results section). If the image is saturated, reduce the laser power.</li>
	<li>Find the brain of the sample by moving the stage and determine its thickness using live view mode in the software by changing the focal planes manually up and down. Set the lower and upper limits of the volume. 
	NOTE: Ensure that the entire brain is contained in the field of view along both the lateral and axial directions.</li>
	<li>Set a z-step size considering the axial resolution of the microscope. 
	NOTE: The optimal z-step size for imaging the larval zebrafish brain depends on the imaging modality and the resolution of the microscope. As an example, a z-step size of 5 &#181;m was used considering the thickness of the light sheet and the average diameter of the cell bodies10.</li>
	<li>Proceed with the image acquisition for the set field of view.
	<ol>
		<li>For volumetric structural imaging, acquire a 3-D image (x,y,z) of the entire brain by changing the focal planes and obtaining 2-D images of each z-plane sequentially.</li>
		<li>For functional imaging of a single z-plane, acquire time-series images (x,y,t) of the neuronal activity of the brain at a certain depth.</li>
		<li>For volumetric functional imaging, acquire a 4-D image (x,y,z,t) of the neuronal activity in the entire brain by obtaining 3-D images sequentially. 
		NOTE: Set the number of frames considering the available memory size of the computer. An acquisition time of less than 1 h is recommended due to the duration of the effect of pancuronium bromide.</li>
	</ol>
	</li>
	<li>After acquiring images, save the results and record the imaging parameters (e.g., pixel size, z-step size, frame rate, laser power) for image analysis.</li>
	<li>Export the images in an appropriate format for data rendering and image analysis. 
	&#8203;NOTE: Exporting images in tagged image file (TIF) format format is recommended. TIF supports lossless image compression and is compatible with most image processing software and programming languages. Additionally, the TIF format supports the inclusion of metadata, such as acquisition parameters, image resolution, and other relevant information that can aid in data interpretation and reproducibility.</li>
</ol>
5. Setup for visualizations using napari
NOTE: napari is an open-source multi-dimensional image viewer in a Python environment with graphics processing unit (GPU)-based rendering33. The napari-animation plugin provides a programmatic creation of movies. Using Fiji, an open-source image processing program, is recommended for general purpose image processing, such as filtering and geometric transformation (see Table of Materials). The source code used for the visualization using napari is available on GitHub (https://github.com/NICALab/Zebrafish-brain-visualization).
<ol>
	<li>Install napari and napari-animation using either pip or conda. After installation, create a new jupyter notebook file. 
	NOTE: Running with jupyter notebook, which is an interactive tool for Python, is recommended over the Python script.</li>
	<li>Import napari and napari-animation.</li>
</ol>
6. Visualizing structures using napari
<ol>
	<li>To visualize images and create movies of the rendered zebrafish brain, load the 3-D image (x,y,z) and open the napari window. Connect the napari-animation plugin (Figure 3A).</li>
	<li>Set parameters such as voxel size, colormap, and contrast limits in the layer controls.</li>
	<li>Adjust the viewer settings (e.g., perspective, angles) in the canvas.</li>
	<li>To capture the rendered image, press the capture button in the animation wizard.</li>
	<li>To generate movies of the rendered volume, modify the viewer settings and add key frames (Figure 3B).</li>
	<li>After adding key frames, set the number of frames (steps) between key frames in the animation wizard. Save the rendered animation.</li>
</ol>
7. Image processing and visualizing neuronal activity using napari
NOTE : To visualize the time-series images of neuronal activity as overlaid images of a static background and activity, a decomposition algorithm must be applied to the raw images. Use a MATLAB implementation of a decomposition algorithm called BEAR24. The MATLAB version of BEAR is available on GitHub (https://github.com/NICALab/BEAR).
<ol>
	<li>To decompose the static background and neuronal activity, apply BEAR to the raw time-series images (x,y,t or x,y,z,t). After the decomposition, save the images of the background and neuronal activity as TIF files (Figure 3C).</li>
	<li>Load the background and neuronal activity images and open the napari window. Connect the napari-animation plugin.</li>
	<li>Use a gray colormap for background images and a hot colormap for neuronal activity images (Figure 3C).</li>
	<li>Set parameters such as opacity and contrast limits in the layer controls.</li>
	<li>To generate movies of neuronal activity, modify the viewer settings and add key frames to render the animation.</li>
	<li>After adding key frames, set the number of frames (steps) between key frames in the animation wizard. Save the rendered animation.</li>
</ol>

In Vivo Whole-Brain Imaging of Zebrafish Larvae

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The authors declare no conflicts of interest.

The current protocol enables in vivo whole-brain imaging of larval zebrafish over an extended period (e.g., longer than 1 h) and visualizations of the acquired structural and functional imaging data.
The most critical steps are sample mounting and positioning. During sample embedding, it is crucial to prevent bubble formation and avoid dust in the agarose gel. If the gel contains air bubbles and dust, the image quality may be severely degraded. While positioning the sample using forceps, it is important to ensure that the sample is level both horizontally and vertically. Otherwise, the regions of interest may not be contained within the field of view of the microscope. In addition, this positioning should be done within a short time window before solidification of the agarose gel to avoid causing damage to its surface, as such damage compromises the image quality.
Another major challenge is to avoid motion in images that come from incomplete solidification of the agarose gel (Figure 6) and paralyzation of the zebrafish. When egg water is added to the sample before complete solidification of the agorse gel, fish move slowly both laterally and axially, manifesting as sample drift in the time-series images (Figure 6A). If the dose of paralytics is insufficient or the duration of effect ends, the sample may attempt to move, which appears as rapid twitching in the time-lapse images.
Despite the importance of the aforementioned steps for acquiring high-quality images without motion artifacts, publicly available protocols15,16,17,18,19 provide only a brief overview of the experimental procedure, lacking these details. For instance, the acquired images without immobilization protocols11 suffer from substantial motion artifacts that make the downstream image analysis challenging. By integrating essential components, such as agarose gel solidification and sample paralyzation, our protocol significantly enhances the consistency in the quality of acquired images while minimizing motion artifacts.
In summary, an optimized and reproducible experimental procedure for in vivo imaging of the larval zebrafish brain is described. The validity and reproducibility of this protocol for in vivo imaging of brain activity and structure have been confirmed in multiple settings14,18,29,30,31. The present workflow focused on the whole-brain imaging of larval zebrafish, but it can be easily applied to imaging other organs of larval zebrafish35,36.

Zebrafish (Danio rerio) has been widely adopted as a model vertebrate animal in neuroscience, owing to its optical transparency at the larval stage, its fast development, its low maintenance cost, and the availability of diverse genetic tools1,2,3,4. In particular, the optical transparency of larvae combined with genetically encoded fluorescent reporters of biological events5,6,7,8,9 provides a unique opportunity to image both neuronal activity and structure at the whole-brain level10,11,12,13,14. However, even with a microscope that supports cellular resolution, the acquired images do not necessarily retain the information at the single-cell level; the optical image quality may be degraded due to the aberration introduced by the agarose gel used for sample mounting, the fish may be mounted at an angle, thus the regions of interest are not fully contained within the field of view of the microscope, and the fish may move during the recording, causing motion artifacts in the images or impeding accurate signal extraction from the images.
Thus, an effective and reproducible protocol is needed to acquire high-quality image data with minimal noise and motion. Unfortunately, publicly available protocols for imaging a whole brain of larval zebrafish in vivo15,16,17,18,19 only briefly describe the procedure, leaving substantial parts of the details, such as agarose solidification, precise mounting techniques, and sample positioning using forceps, up to each experimentalist. In addition, inconsistencies in agarose concentration and immobilization methods10,11,14,15,16,17,18,19 can lead to challenges arising from fish movement during the imaging process.
Here, a detailed protocol for whole-brain imaging of larval zebrafish using three-dimensional fluorescence microscopy is provided. Figure 1 provides a graphical overview of the protocol: sample preparation and immobilization, sample embedding, image acquisition, and visualization after imaging. The structural and functional imaging of the larval zebrafish brain in vivo is demonstrated using a commercial confocal microscope and a custom-designed fluorescence microscope. This protocol can be tailored by practitioners for brain imaging with certain sensory stimuli or behavior contexts depending on the experimental needs and design.

<table><tbody><tr><td>1.5 mL microcentrifuge tube</td><td>SciLab</td><td>SL.Tub3513</td><td>To aliquot agarose gel and pancuronium bromide solution</td></tr><tr><td>15 mL Falcon tubes</td><td>Falcon</td><td>352096</td><td>To prepare agarose gel and pancuronium bromide solution</td></tr><tr><td>16&amp;times; 0.8NA water dipping objective lens</td><td>Nikon</td><td>CFI75 LWD 16&amp;times;W</td><td>Objective lens for whole-brain imaging</td></tr><tr><td>1-phenyl 2-thiourea (PTU)</td><td>Sigma-Aldrich</td><td>P7629-10G</td><td>200 &amp;mu;M of 1-phenyl 2-thiourea (PTU)</td></tr><tr><td>50 mL Falcon tubes</td><td>Falcon</td><td>352070</td><td>To prepare egg water</td></tr><tr><td>Disposable transfer pipette</td><td>SciLab</td><td>SL.Pip3032</td><td>To transfer zebrafish larvae</td></tr><tr><td>Egg water</td><td>N/A</td><td>N/A</td><td>0.6 g sea salt in 10 L deionized water</td></tr><tr><td>Forceps</td><td>Karl Hammacher GmbH</td><td>HSO 010-10</td><td>Forceps used for sample positioning</td></tr><tr><td>Low melting point agarose</td><td>Thermo Scientific</td><td>R0801</td><td>2% (wt/vol) agarose gel</td></tr><tr><td>Napari</td><td>Napari</td><td>N/A</td><td>To visualize microscopy images in 3-D</td></tr><tr><td>NIS-Elements C</td><td>Nikon</td><td>N/A</td><td>Imaging software for confocal microscope</td></tr><tr><td>Pancuronium bromide</td><td>Sigma-Aldrich</td><td>P1918-10MG</td><td>0.25 mg/mL of pancuronium bromide solution</td></tr><tr><td>Petri dish, 35 mm</td><td>SPL Life Sciences</td><td>11035</td><td>Petri dish used for sample embedding</td></tr><tr><td>Petri dish, 55 mm</td><td>SPL Life Sciences</td><td>11050</td><td>To prepare zebrafish larvae after screening</td></tr><tr><td>Point-scanning confocal microscopy system (C2 Plus)</td><td>Nikon</td><td>N/A</td><td>Confocal microscope for whole-brain imaging</td></tr><tr><td>Sea salt&amp;nbsp;</td><td>Sigma-Aldrich</td><td>S9883-500G</td><td>Sea salt used for preparing egg water</td></tr></tbody></table>

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.

The structure and neuronal activity of the larval zebrafish brains expressing calcium indicator pan-neuronal GCaMP7a (Tg(huc:GAL4);Tg(UAS:GCaMP7a))20,21,22with a casper (mitfa(w2/w2);mpv17(a9/a9))34 background was imaged at 3-4 d.p.f. by following the described protocol.
For volumetric structural imaging, the specimen was imaged using a commercial point-scanning confocal microscopy system equipped with a 16x 0.8 NA water dipping objective lens. A 488 nm excitation laser was used for both structural and functional imaging. The frame rate, image resolution, pixel size, and axial step size were 0.25 Hz, 2048 x 2048, 0.34 &#181;m, and 1.225 &#181;m, respectively. The image acquisition took approximately 1 h and 20 min. The volumetric field of view of the acquired image covered the brain regions of the forebrain, midbrain, and hindbrain (Figure 4A). Neuronal cell bodies of the medulla oblongata, cerebellar plate, optic tectum, habenula, and dorsal telencephalon in the whole brain of 4 d.p.f. larval zebrafish were clearly visible in the confocal microscopy images (Figure 4B). The 3-D rendering of the confocal microscopy images was performed using napari27 by following the aforementioned protocol (Figure 4C and Video 1).
For functional imaging in 2-D, the specimen was imaged using the same confocal microscopy system, equipped with a 16x 0.8 NA water dipping objective lens. The frame rate, image resolution, and pixel size were 2 Hz, 512 x 256, and 1.5 &#181;m, respectively. The neuronal cell bodies were clearly visible in both the background and the superimposed neuronal activity (Figure 5A,B).
For 3-D functional imaging, a custom-designed 3-D microscopy system18 was used, which was capable of in vivo imaging the neuronal activity of a whole larval zebrafish brain with a field of view of 1,040 &#181;m &#215; 400 &#181;m &#215; 235 &#181;m and lateral and axial resolutions of 1.7 &#181;m and 5.4 &#181;m, respectively. The imaging rate was up to 4.2 volumes per second. Similar to rendering the structural imaging data, napari was used for 3-D rendering of the whole-brain calcium imaging data (Figure 5C and Video 2).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65218/65218fig01.jpg" /> 
Figure 1: Overview of the experimental procedure. Zebrafish sample preparation and paralyzation (step 1). The 2% (wt/vol) agarose gel preparation (step 2). Sample mounting and positioning (step 3). Image acquisition (step 4). Image processing and visualizing structure and neuronal activity (step 5-7). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65218/65218fig01large.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/65218/65218fig02.jpg" /> 
Figure 2: Experimental procedure for preparing whole-brain imaging. (A) Screened zebrafish sample expressing pan-neuronal GCaMP7a in a Petri dish filled with egg water. (B) Equipment and material required for sample mounting and positioning. (1) Heat block at 37 &#176;C; (2) 2% (wt/vol) agarose gel; (3) 1.5 mL microcentrifuge tube; (4) egg water; (5) 0.25 mg/mL pancuronium bromide solution; (6) Petri dish; (7) forceps; (8) transfer pipette. (C) Stereomicroscope image of the paralyzed sample. The black arrow points to the heart of the sample. (D) The sample in the 1.5 mL microcentrifuge tube is transferred using a pipette. The inset shows an enlarged view of the boxed area. (E) The sample (black arrow) is placed in the center of the Petri dish using forceps. (F) The sample is aligned using forceps. (G) An example of a misaligned embedded sample. (H) An example of an aligned embedded sample. (I) The specimen is placed on a microscope stage under the objective lens for image acquisition. Scale bar: 500 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65218/65218fig02large.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/65218/65218fig03.jpg" /> 
Figure 3: Visualization of larval zebrafish brain images. (A) The 3-D rendering of a confocal microscopy image of a larval zebrafish (4 d.p.f) brain expressing pan-neuronal GCaMP7a. napari, an open-source multi-dimensional image viewer in a Python environment, was used for the rendering. The napari window comprises layer controls (red box), a layer list (yellow box), a canvas (blue box), and an animation wizard (green box). (B) Key frames with multiple viewer settings are added for rendering an animation. (C) Confocal microscopy image of 2-D time-lapse imaging of the neuronal activity in the larval zebrafish brain. The image is decomposed to the background (left) and neuronal activity (middle), then overlaid (right). Scale bar: 100 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65218/65218fig03large.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/65218/65218fig04.jpg" /> 
Figure 4: Imaging structure of a larval zebrafish brain. (A) Maximum intensity projection (MIP) of a confocal microscopy image of a larval zebrafish (4 d.p.f.) brain expressing pan-neuronal GCaMP7a. Top: lateral MIP. Bottom: axial MIP. Each boundary contains the forebrain (red), midbrain (yellow), and hindbrain (blue). (B) A total of 10 axial slices at multiple depths from a volumetric image of the brain (at z = 25 &#181;m, 50 &#181;m, 75 &#181;m, 100 &#181;m, 125 &#181;m, 150 &#181;m, 175 &#181;m, 200 &#181;m, 225 &#181;m, 250 &#181;m, counted upward from top to bottom; z = 0 &#181;m indicates the top surface of the brain). Each white box represents the region of the brain (medulla oblongata, cerebellar plate, optic tectum, habenula, and dorsal telencephalon). (C) The whole brain rendered using napari.Scale bar: 100 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65218/65218fig04large.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/65218/65218fig05.jpg" /> 
Figure 5: Imaging neuronal activity of a larval zebrafish brain. (A) Confocal microscopy image of neuronal activity in a larval zebrafish (4 d.p.f.) brain expressing pan-neuronal GCaMP7a. Scale bar: 100 &#181;m. (B) Enlarged view of the boxed area in A showing the neuronal activity in the optic tectum at multiple time points. Scale bar: 20 &#181;m. (C) The 3-D rendering of the whole-brain neuronal activity in the larval zebrafish brain acquired using a custom-designed microscope (left: t = 133 s; right: t = 901 s). The neuronal activity is superimposed on the static background. Scale bar: 100 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65218/65218fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/65218/65218fig06.jpg" /> 
Figure 6: Time-lapse imaging with and without sample drift. (A) Time-lapse imaging of the larval zebrafish brain expressing pan-neuronal GCaMP7a with sample drift. Egg water was added to the sample before solidification of the agarose gel (step 3.6-3.7). The fish moved in the lateral and axial directions. (B) Time-lapse imaging of a larval zebrafish brain without sample drift. Egg water was added to the sample after solidification of the agarose gel (step 3.6-3.7). Scale bar: 100 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65218/65218fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Video 1: 3-D rendering of the whole-brain structure of a larval zebrafish (4 d.p.f.) brain expressing pan-neuronal GCaMP7a. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65218/Video1.zip" target="_blank">Please click here to download this Video.</a>
Video 2: 3-D rendering of whole-brain neuronal activity in a larval zebrafish (4 d.p.f.) brain expressing pan-neuronal GCaMP7a. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65218/Video2.zip" target="_blank">Please click here to download this Video.</a>

Watch this Scientific Journal Video about In Vivo Whole-Brain Imaging of Zebrafish Larvae Using Three-Dimensional Fluorescence Microscopy at JoVE.com

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.

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 ...

in-vivo-whole-brain-imaging-zebrafish-larvae-using-three-dimensional

Nanyang Technological University

Research

JoVE Journal

Neuroscience

4.1K Views.  KAIST. We ontwikkelen optische immuunsystemen en computationele algoritmen om de neuroactiviteit van de hele hersenen met een hoge ruimtelijke en temporele resolutie vast te leggen en te analyseren. Proefvis is een ideaal modeldier voor onderzoek. Dankzij elke optische transparantie en de beschikbaarheid van diverse genetische hulpmiddelen De optische beeldkwaliteit kan worden aangetast als gevolg van de aberratie die wordt geïntroduceerd door de Agarose-gel die wordt gebruikt voor monstermontage, ...

Video: Auteur in de schijnwerpers: in vivo beeldvorming van de hele hersenen van zebravislarven met behulp van driedimensionale fluorescentiemicroscopie  

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

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

In recent years, the zebrafish embryo has emerged as a popular model to study developmental biology due to traits such as ex utero embryo development and optical transparency. In particular, the zebrafish embryo has become an important organism to study vertebrate kidney organogenesis as well as multiciliated cell (MCC) development. To visualize MCCs in the embryonic zebrafish kidney, we have developed a combined protocol of whole-mount fluorescent in situ hybridization (FISH) and whole mount immunofluorescence (IF) that enables high resolution imaging. This manuscript describes our technique for co-localizing RNA transcripts and protein as a tool to better understand the regulation of developmental programs through the expression of various lineage factors.

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

Cilia development is vital to proper organogenesis. This protocol describes an optimized method to label and visualize ciliated cells of the zebrafish.

In recent years, the zebrafish embryo has emerged as a popular model to study developmental biology due to traits such as ex utero embryo development and ...

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

A Layered Mounting Method for Extended Time-Lapse Confocal Microscopy of Whole Zebrafish Embryos

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues or cells. A difficulty with imaging whole embryo development is that zebrafish embryos grow substantially in length. When mounted as regularly done in 0.3-1% low melt agarose, the agarose imposes growth restriction, leading to distortions in the soft embryo body. Yet, to perform confocal time-lapse microscopy, the embryo must be immobilized. This article describes a layered mounting method for zebrafish embryos that restrict the motility of the embryos while allowing for the unrestricted growth. The mounting is performed in layers of agarose at different concentrations. To demonstrate the usability of this method, whole embryo vascular, neuronal and muscle development was imaged in transgenic fish for 55 consecutive hours. This mounting method can be used for easy, low-cost imaging of whole zebrafish embryos using inverted microscopes without requirements of molds or special equipment.

This article describes a method to mount fragile zebrafish embryos for extended time-lapse confocal microscopy. This low-cost method is easy to perform using regular glass-bottom microscopy dishes for imaging on any inverted microscope. The mounting is performed in layers of agarose at different concentrations.

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues ...

Visualizing the Developing Brain in Living Zebrafish using Brainbow and Time-lapse Confocal Imaging

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in vivo imaging techniques can provide a clear window into these processes in the living organism. For example, dividing cells and their progeny can be followed in real time as the nervous system forms. In recent years, technical advances in multicolor techniques have expanded the types of questions that can be investigated. The multicolor Brainbow approach can be used to not only distinguish among like cells, but also to color-code multiple different clones of related cells that each derive from one progenitor cell. This allows for a multiplex lineage analysis of many different clones and their behaviors simultaneously during development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells over real time within the developing zebrafish nervous system. This is particularly useful for following cellular interactions among like cells, which are difficult to label differentially using traditional promoter-driven colors. Our approach can be used for tracking lineage relationships among multiple different clones simultaneously. The large datasets generated using this technique provide rich information that can be compared quantitatively across genetic or pharmacological manipulations. Ultimately the results generated can help to answer systematic questions about how the nervous system develops.

In vivo imaging is a powerful tool that can be used to investigate the cellular mechanisms underlying nervous system development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells in real time within the developing zebrafish nervous system.

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in ...

Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish

Zebrafish are a powerful tool to study developmental biology and pathology in vivo. The small size and relative transparency of zebrafish embryos make them particularly useful for the visual examination of processes such as heart and vascular development. In several recent studies transgenic zebrafish that express EGFP in vascular endothelial cells were used to image and analyze complex vascular networks in the brain and retina, using confocal microscopy. Descriptions are provided to prepare, treat and image zebrafish embryos that express enhanced green fluorescent protein (EGFP), and then generate comprehensive 3D renderings of the cerebrovascular system. Protocols include the treatment of embryos, confocal imaging, and fixation protocols that preserve EGFP fluorescence. Further, useful tips on obtaining high-quality images of cerebrovascular structures, such as removal the eye without damaging nearby neural tissue are provided. Potential pitfalls with confocal imaging are discussed, along with the steps necessary to generate 3D reconstructions from confocal image stacks using freely available open source software.

Imaging of cerebrovascular development in larval zebrafish is described. Techniques to facilitate 3D imaging and modify cerebrovascular development using chemical treatments are also provided.

Zebrafish are a powerful tool to study developmental biology and pathology in vivo.  The small size and relative transparency of zebrafish embryos make ...

Analysis of Zebrafish Cardiac Contraction with Virtual Reality Interaction

4D Light-sheet Imaging of Zebrafish Cardiac Contraction

Zebrafish is an intriguing model organism known for its remarkable cardiac regeneration capacity. Studying the contracting heart in vivo is essential for gaining insights into structural and functional changes in response to injuries. However, obtaining high-resolution and high-speed 4-dimensional (4D, 3D spatial + 1D temporal) images of the zebrafish heart to assess cardiac architecture and contractility remains challenging. In this context, an in-house light-sheet microscope (LSM) and customized computational analysis are&#160;used to overcome these technical limitations. This strategy, involving LSM system construction, retrospective synchronization, single cell tracking, and user-directed analysis, enables one to investigate the micro-structure and contractile function across the entire heart at the single-cell resolution in the transgenic Tg(myl7:nucGFP) zebrafish larvae. Additionally, we are able to further incorporate microinjection of small molecule compounds to induce cardiac injury in a precise and controlled manner. Overall, this framework allows one to track physiological and pathophysiological changes, as well as the regional mechanics at the single-cell level during cardiac morphogenesis and regeneration.

Author Spotlight: High-Resolution 4D Light-Sheet Imaging and Virtual Reality in Zebrafish for Single-Cell Analysis of Heart Function

This protocol utilizes light-sheet imaging to investigate cardiac contractile function in zebrafish larvae and gain insights into cardiac mechanics through cell tracking and interactive analysis.

Zebrafish is an intriguing model organism known for its remarkable cardiac regeneration capacity. Studying the contracting heart in vivo is essential for ...

Bioengineering

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical clarity and amenability to genetic manipulation, the zebrafish embryo has become a popular model organism with which to perform time-lapse analysis of morphogenesis in living embryos. Confocal imaging of a live zebrafish embryo requires that a tissue of interest is persistently labeled with a fluorescent marker, such as a transgene or injected dye. The process demands that the embryo is anesthetized and held in place in such a way that healthy development proceeds normally. Parameters for imaging must be set to account for three-dimensional growth and to balance the demands of resolving individual cells while getting quick snapshots of development. Our results demonstrate the ability to perform long-term in vivo imaging of fluorescence-labeled zebrafish embryos and to detect varied tissue behaviors in the cranial neural crest that cause craniofacial abnormalities. Developmental delays caused by anesthesia and mounting are minimal, and embryos are unharmed by the process. Time-lapse imaged embryos can be returned to liquid medium and subsequently imaged or fixed at later points in development. With an increasing abundance of transgenic zebrafish lines and well-characterized fate mapping and transplantation techniques, imaging any desired tissue is possible. As such, time-lapse in vivo imaging combines powerfully with zebrafish genetic methods, including analyses of mutant and microinjected embryos.

Time-lapse confocal imaging is a powerful technique useful for characterizing embryonic development. Here, we describe the methodology and characterize craniofacial morphogenesis in wild-type, as well as pdgfra, smad5, and smo mutant embryos.

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical ...

4-Dimensional Imaging of Zebrafish Optic Cup Morphogenesis

Visual system function requires the establishment of precise tissue and organ structures. In the vertebrate eye, structural defects are a common cause of visual impairment, yet mechanisms of eye morphogenesis are still poorly understood. The basic organization of the embryonic eye is conserved throughout vertebrates, thus live imaging of zebrafish embryos has become a powerful approach to directly observe eye development at real time under normal and pathological conditions. Dynamic cell processes including movements, morphologies, interactions, division, and death can be visualized in the embryo. We have developed methods for uniform labeling of subcellular structures and timelapse confocal microscopy of early eye development in zebrafish. This protocol outlines the method of generating capped mRNA for injection into the 1-cell zebrafish embryo, mounting embryos at optic vesicle stage (~12 hours post fertilization, hpf), and performing multi-dimensional timelapse imaging of optic cup morphogenesis on a laser scanning confocal microscope, such that multiple datasets are acquired sequentially in the same imaging session. Such an approach yields data that can be used for a variety of purposes, including cell tracking, volume measurements, three-dimensional (3D) rendering, and visualization. Our approaches allow us to pinpoint the cellular and molecular mechanisms driving optic cup development, in both wild type and genetic mutant conditions. These methods can be employed directly by other groups or adapted to visualize many additional aspects of zebrafish eye development.

This protocol describes an approach for in toto labeling and multidimensional imaging of zebrafish early eye development. We describe labeling, embedding, and four dimensional (4D) imaging using laser scanning confocal microscopy, and considerations for optimizing acquisition of datasets for dissecting mechanisms of optic cup morphogenesis.

Visual system function requires the establishment of precise tissue and organ structures. In the vertebrate eye, structural defects are a common cause of ...

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 ...

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

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