We thank Y. A. Pan, J. Livet and Z. Tobias for technical and intellectual contributions. This work was supported by the National Science Foundation (Award 1553764) and the M.J. Murdock Charitable Trust.

Procedures involving animal subjects have been approved by the Institutional Animal Care and Use Committee (IACUC) at Lewis &#38; Clark College.
1. Microinjection of Zebrafish Embryos
<ol>
	<li>Set up wild type, adult zebrafish in sex-segregated mating tanks the afternoon prior to performing microinjections39,40.</li>
	<li>Prepare the DNA solution in the morning of the microinjections. Dilute hsp:Zebrabow11 plasmid DNA to a concentration of ~10 ng/&#956;L in 0.1 mM KCl, along with 2.5% phenol red and 3.75U Cre recombinase enzyme.</li>
	<li>Perform microinjections of DNA solution into one-cell zebrafish embryos within 45 min of fertilization39,41. Inject approximately 4.2 nL of this solution into each embryo, equivalent to ~42 pg of plasmid DNA. 
	NOTE: The microinjection step can be omitted if a transgenic, Brainbow-expressing zebrafish line is mated instead, such as Tg(ubi:Zebrabow)15 and Tg(neurod:Zebrabow)18. Performing microinjections can be advantageous because it allows researchers to titrate the copy number and labeling density. Furthermore, a second DNA construct that tags or manipulates a specific gene product can be injected alongside Brainbow if desired (e.g., a far-red fluorescent protein that complements Brainbow18).</li>
	<li>Maintain injected embryos in Petri dishes of E3 medium39 in a 28 &#176;C incubator for 24 h.</li>
</ol>
2. Heat Shock to Induce Brainbow Expression
NOTE: If the plasmid DNA injected or the transgene expressed does not utilize the hsp70 promoter to drive the expression, the heat shock step can be skipped, and healthy embryos should be immediately transferred to phenylthiourea (PTU) at 24 h post fertilization (hpf).
<ol>
	<li>At 24 hpf, cull dead and deformed embryos from the group of injected embryos. Then, transfer the healthy embryos to a 50 mL tube, with up to 20 embryos/tube.</li>
	<li>Fill the tubes with 10 mL of E3. Place a cap on top of each tube but do not close tightly.</li>
	<li>Place the tube rack containing the 50 mL tubes upright in a 37 &#176;C water bath. Ensure that the water level in the bath is higher than the level of the E3 in the tubes and leave them for 80 to 90 min.</li>
	<li>Remove the tube rack with the embryos from the water bath and return to the 28 &#176;C incubator upright. Allow up to 1 h for the E3 to cool and the embryos to gradually re-acclimate to the temperature. Then, transfer the embryos to Petri dishes of 0.2 mM PTU in E3 warmed in the incubator to prevent pigmentation of embryos. 
	CAUTION: PTU is a toxic chemical that should be handled with care and disposed of appropriately. Wearing gloves is suggested.</li>
</ol>
3. Screening Embryos for Brainbow Expression
<ol>
	<li>At 2 to 4 h after the heat shock, examine the embryos under a standard fluorescence dissection microscope for expression of CFP or YFP, which indicates successful Brainbow recombination (from default expression of dTomato). 
	NOTE: If multiple fluorescence filter options are available, screening for CFP is the most efficient way to identify well-recombined fish. If CFP and YFP filters are not available, YFP can also be dimly visualized under green fluorescent protein (GFP) filters due to the overlap in their spectral profiles.</li>
	<li>Select embryos with robust FP expression throughout and transfer to a separate dish with PTU. Image embryos at 1 day post fertilization (dpf; 28 hpf or later) or maintain embryos in PTU and image at 2 or 3 dpf.</li>
</ol>
4. Mounting Embryos for In Vivo Imaging
<ol>
	<li>Prior to the day of the experiment, prepare the imaging chamber and embryo manipulation tool.
	<ol>
		<li>Prepare the imaging chamber by carefully supergluing a plastic ring to the center of a 60 mm Petri dish.</li>
		<li>Optionally, prepare a custom-made embryo manipulator to move embryos within the dish and orient embryos while mounting. Construct the manipulator by supergluing a small length of nylon fishing line (~1/2 in, ~6 lb) to the end of a wooden cotton swab stick that has been cut to a ~4 in length.</li>
	</ol>
	</li>
	<li>If necessary (i.e., imaging at 2 dpf or earlier), before mounting the embryos dechorionate using syringes under a dissection microscope.</li>
	<li>Anesthetize zebrafish embryos.
	<ol>
		<li>Fill a 60 mm Petri dish halfway with E3 and add 5&#8211;6 drops of 4 mg/mL MS-222 Tricaine-S to a final concentration of ~0.2 mM. Swirl to mix. 
		CAUTION: Tricaine should be handled with care and disposed of appropriately.</li>
		<li>Transfer embryos from the PTU to the tricaine solution, ensuring as little PTU is transferred as possible. While fish reflexes should cease after 1&#8211;2 min, leave the embryos in tricaine for up to 10 min to ensure thorough anesthetization for time-lapse imaging experiments.</li>
		<li>Confirm that the embryos are properly anesthetized. Assess the startle reflex by touching the embryo tails gently with the embryo manipulator. If a reflex movement is observed, add an additional drop of tricaine to the dish as far from the embryos as possible and swirl to mix. Observe the embryo heart rate under a dissection scope. If it is abnormally slow, add additional drops of E3 to the dish to reduce the tricaine concentration.</li>
	</ol>
	</li>
	<li>Mount anesthetized embryos in agarose.
	<ol>
		<li>Transfer the fish to the center of plastic ring within the imaging chamber and remove as much excess E3 as possible using a fine-tipped transfer pipette.</li>
		<li>Using a clean transfer pipette, cover the fish with 1.0% low-melt agarose (LMA) in E3 stored at 42 &#176;C and fill the entire plastic ring with a thin layer of agarose. Use a transfer pipette to gently pull the embryos up into the pipette tip and back into the agarose without introducing air bubbles.</li>
		<li>Before the agarose hardens, quickly use an embryo manipulator to orient the fish appropriately. If imaging with an upright microscope, position the embryos as close to the upper surface of the agarose as possible. Position the embryos parallel to the bottom of the imaging chamber with the tail straight. 
		NOTE: For hindbrain imaging, the embryos can be positioned in a dorsal or sagittal view. Care should be taken to ensure that the heads of the embryos do not sink while the agarose is still liquid. Other orientations may be appropriate to target other developing tissues for imaging. For inverted microscopes, mounting would be done differently, typically positioning the fish close to the bottom of a glass-bottomed Petri dish.</li>
	</ol>
	</li>
	<li>Wait for the agarose to harden, then fill the imaging chamber with E3. Add as much E3 as possible to account for the evaporation over the course of imaging.</li>
</ol>
5. Time-lapse Confocal Imaging of Developing Zebrafish Hindbrain
<ol>
	<li>Place the imaging chamber on the confocal microscope in preparation for the imaging.
	<ol>
		<li>Place the imaging chamber with the fish and E3 on the confocal microscope.</li>
		<li>Select an objective with a high numerical aperture and a long working distance, such as a 20x water immersion objective (1.0 NA).</li>
		<li>Find a region with appropriately dense and bright labeling of the cell type(s) of interest. 
		NOTE: A temperature-controlled stage/apparatus on the microscope may also be used to regulate temperature and humidity during long imaging sessions. This can decrease evaporation.</li>
	</ol>
	</li>
	<li>Set up the acquisition parameters for Brainbow imaging.
	<ol>
		<li>Image each FP channel sequentially. If using a commercial software (e.g., Zen software), this is done by preparing three &#8220;tracks&#8221;. Use an Argon laser to excite CFP at 458 nm and YFP at 514 nm. Use a DPSS 561 nm laser to excite dTomato. Collect emissions between 463&#8211;509 nm for CFP, 519&#8211;555 nm for YFP, and 566&#8211;691 nm for dTomato.</li>
		<li>For on-screen display, code CFP as blue, YFP as yellow, and dTomato as red. 
		NOTE: These settings will vary depending upon which laser lines and other features are available on the confocal microscope. To increase the speed of imaging, CFP and dTomato channels can be imaged simultaneously without appreciable bleed-through. If a separate FP is also being imaged, such as a far-red FP, this can be imaged sequentially or simultaneously with YFP.</li>
		<li>Set up the general imaging parameters to take 16-bit images with a resolution of at least 1024 x 1024 and averaging twice. Adjust the zoom depending on the region and cell type of interest (i.e., for hindbrain imaging with a 20x objective the zoom will range from 1.0&#8211;2.5). Maximize the field of view to allow for growth of the fish over time.</li>
		<li>Viewing one track at a time (and turning off other lasers to prevent photobleaching), optimize the acquisition settings for each FP individually (e.g., laser power, pinhole size, photomultiplier tube gain, etc.).</li>
		<li>Select Z-stack range to image. 
		NOTE: For long imaging sessions (&#62;2 h), take into consideration that the fish will continue to grow throughout the course of imaging, thus both the XY field and the Z-stack range should account for this with extra room. If imaging the hindbrain, include space in the field for the tissue to move rostrally during the imaging period. Space also needs to be included for growth in the Z dimension. Include in the image approximately 10&#8211;20 &#956;m above the region of interest, whether the fish is positioned for sagittal or dorsal view.</li>
	</ol>
	</li>
	<li>Set up parameters for time-lapse imaging.
	<ol>
		<li>Select a time interval. Interval length ranges between 10&#8211;30 min to track mitotic and apoptotic events in the developing hindbrain. Interval length will vary depending on the speed of the events of interest and the total time it takes to capture a single Z-stack.</li>
		<li>Select the length of the imaging session. Time-lapse imaging sessions generally occur overnight and can last at least 16 h.</li>
	</ol>
	</li>
	<li>Run the experiment. 
	NOTE: The fish can be euthanized immediately following imaging or carefully dissected from the agarose using syringes and returned to E3 to recover. Recovered fish can then be imaged again at a later time point, though cell position and depth will shift during this period due to overall growth.</li>
</ol>
6. Time-lapse File Management
<ol>
	<li>Import the image file into the analysis software.
	<ol>
		<li>If using Zen software, save the file in .czi format once image acquisition is completed. Make sure to save raw data in a format compatible with Fiji42 and/or other software.</li>
		<li>Import into Fiji software using BioFormats Importer. 
		NOTE: Time-lapse files will be large and can require significant amounts of time and memory to open for analysis. Thus, it is helpful to be strategic in how they are saved and opened. It can be useful to save a lower-quality version that can be opened more easily to search for events of interest by eye.</li>
	</ol>
	</li>
	<li>If using Fiji, create smaller, more manageable subsets of time-lapse files. Generate subsets either by time (e.g., viewing full Z-stack at the first time point) or by depth (e.g., viewing the first 50 Z-sections at each time point) by clicking Image|Stacks|Tools|Make Substack.</li>
</ol>
7. Quantitative Clonal Color Analysis of Brainbow Images
<ol>
	<li>Measure the mean red, green, and blue (RGB) intensity values for the cells of interest in Fiji. 
	NOTE: These instructions are specific to image analysis in Fiji. However, researchers may prefer to obtain mean RGB intensity values in an alternative software program.
	<ol>
		<li>Ensure that the file is correctly cropped so that there is no black space around the edges of the image. Black space will artificially lower the minimal intensity measurements needed for the analysis. If the file does need to be cropped, select the Rectangle Selection Button and draw a region of interest (ROI) around the field of view, excluding all black space. Click Image|Crop.</li>
		<li>Set the measurement tool to take mean, minimum, and maximum gray value measurements. Click Analyze|Set Measurements. Ensure the checkboxes for Min &#38; Max Gray Value and Mean Gray Value are both selected.</li>
		<li>Viewing the raw data files or subsetted Z-stacks in Fiji, find cells of interest for clonal color analysis. In the hindbrain, putative clones can be visually identified by their shared hue as well as their radial orientation. Do not select cells that are in close contact with, and thus difficult to resolve from, neighboring cells of another color. It may be difficult to quantify their hue.</li>
		<li>Find the center Z-plane of the cell of interest using the Z-scrollbar. Right-click the Oval Selection Button and select the Elliptical Selection Button. Other selection tools will be appropriate for different cell types. Draw an elliptical ROI around the central ~2/3 of the cell body.</li>
		<li>Using the C-scrollbar, select the red (dTomato) channel. Take the mean intensity measurement for this channel by selecting Analyze|Measure. Repeat with the same ROI and focal plane for the green (YFP) and blue (CFP) channels and save all mean intensity values.</li>
		<li>Click anywhere on the image to remove the elliptical ROI in the cell of interest.</li>
		<li>To measure background levels for normalization, use the C-scrollbar and select the red (dTomato) channel. Take the minimum intensity measurement for the whole field in this channel by selecting Analyze|Measure. Repeat with the same focal plane for the green (YFP) and blue (CFP) channels and save all minimum intensity values.</li>
	</ol>
	</li>
	<li>Calculate relative RGB channel weights for the cells of interest. 
	NOTE: Calculation of relative RGB channel weights from these intensity values can be performed in a variety of software programs, such as Microsoft Excel or R43.
	<ol>
		<li>Normalize the mean intensity measurement of the cell ROI for each channel by subtracting the minimal intensity from the entire field in that channel and focal plane.</li>
		<li>Sum the normalized mean RGB intensity values from each channel to find the total normalized RGB intensity for the cell.</li>
		<li>Calculate the relative channel weight for each channel by dividing the normalized mean intensity value for that channel by the total normalized RGB intensity.</li>
	</ol>
	</li>
	<li>Display relative RGB channel weights as ternary plots using the Ternary package for R43,44. Ternary plots are useful to visualize the clustering of similar colors in 3D RGB space.</li>
	<li>Calculate the color spread coefficient to quantify the difference between cell colors.
	<ol>
		<li>Calculate the Euclidean distance between the two colors in 3D RGB space using the following equation: 
		<img alt="Equation 1" src="/files/ftp_upload/60593/60593equ01.jpg" /> 
		where R, G, and B are the relative RGB channel weights calculated in 7.2.</li>
		<li>For ease of comprehension, normalize the color distance by dividing by &#8730;2, the maximum possible distance between colors, to obtain a color spread coefficient between 0 and 1, where 0 indicates exactly the same color and 1 indicates completely different colors (e.g., pure red and pure blue).</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

This protocol describes a method to visualize clones of progenitor cells and neurons in the developing zebrafish hindbrain and follow them in vivo using Brainbow and time-lapse confocal microscopy11. The major advantage of this protocol in comparison to in vitro or ex vivo studies is the ability to directly observe the proliferative zone of the vertebrate brain in its natural milieu over time. This technique builds on previous studies&#160;that labeled a single clone using retroviral vectors. In c...

In the early phases of development, pools of specialized progenitor cells divide repeatedly in proliferative zones, producing diverse arrays of daughter cells. The cells born during this developmental period will then differentiate and travel to form the nascent organs. In the nervous system, progenitors such as radial glia give rise to immature neurons in ventricular zones. As neurons migrate away from ventricles and mature, the expanding tissue eventually forms the highly complex structures of the brain1,2,3,4,5,6. The coordination between division of progenitors and differentiation and migration of neurons will determine the eventual size, shape, and thus function of the brain, directly impacting behavior7,8,9,10. While tight control over these processes is clearly crucial for normal brain development, the global mechanisms that regulate these dynamics are not well understood. Here we describe a tool to study nervous system development at a cellular resolution, allowing researchers to visualize progenitor cells and neurons in vivo in the developing zebrafish brain with Brainbow and track their behavior over time via time-lapse confocal microscopy11. The approach can also be adapted to visualize other parts of the developing embryo.
To observe and distinguish among cells in the developing zebrafish brain, we have adapted the Brainbow cell-labeling technique11. Brainbow utilizes the randomly determined, combinatorial expression of three distinct fluorescent proteins (FPs) to label a population of cells. While the default expression for Brainbow expression is the red FP dTomato, recombination by the enzyme Cre recombinase results in expression of mCerulean (cyan fluorescent protein, CFP) or yellow fluorescent protein (YFP)12,13. The combined amount of each FP expressed in a cell gives it a unique hue, allowing clear visual distinction from neighboring cells. Additionally, when a progenitor cell divides, each daughter cell will inherit the color from its mother cell, producing color-coded clones and allowing researchers to trace cell lineage11,14. While originally used to analyze neuronal circuitry in mice12, Brainbow has since been expressed in a wide variety of model organisms, including zebrafish15.
Our technique builds on previous multicolor labeling and imaging methods to directly image multiple color-coded clones over time in living zebrafish. Due to their optical transparency as embryos, zebrafish are well suited to imaging experiments16, and previous studies have utilized Brainbow in zebrafish to study a variety of tissues, including the nervous system11,15,17,18,19,20,21,22,23,24,25,26,27. The ability to directly image into the living organism, along with their rapid ex utero development, make zebrafish a valuable model of vertebrate development. In contrast to the mammalian brain, the entire proliferative zone of the zebrafish hindbrain is readily available for imaging without disruption to its endogenous environment6. This allows experiments to be conducted in the living organism, rather than in in vitro or fixed tissue preparations. In contrast to fixed imaging experiments, in vivo studies allow for a longitudinal design, producing hours of data that can be analyzed for patterns, thus increasing the likelihood of observing relatively rare events. Depending upon the speed and length of the events of interest, researchers may choose to perform short (1&#8211;2 h) or long (up to ~16 h) time-lapse imaging experiments. By using the zebrafish heat shock promoter 70 (hsp70, hsp), Brainbow expression can be temporally controlled28,29. Additionally, the mosaic expression induced by this promoter is well suited for labeling and tracking many clones11.
The ability to visually identify multiple clones within the living brain is an advantage of this method. Important previous studies that investigated the role of clones within development of the nervous system utilized retroviral vectors to label a single progenitor cell and its progeny using a single FP or other readily visualized protein. Such labeling allows researchers to observe a single clone over time, either in vitro or in vivo2,30,31,32,33,34,35,36,37,38. In contrast to methods to track the behavior of cells within one clone, the distinct colors of Brainbow allow researchers to observe dynamics among clones. Additionally, by using Brainbow to label many clones within the brain, additional data on clonal behavior is collected relative to techniques that label a single clone11. Importantly, the approaches described here can be expanded to generate developmental comparisons between fish that have undergone different genetic or pharmacological manipulations18. Overall, these advantages make time-lapse in vivo confocal imaging of Brainbow-expressing zebrafish ideal for researchers exploring development of the vertebrate nervous system, particularly those interested in the role of clones.

<table><tbody><tr><td>1.5 mL transfer pipette (fine tip)</td><td>Globe Scientific, Inc.</td><td>134020</td><td></td></tr><tr><td>1-phenyl-2-thiourea (PTU)</td><td>Alfa Aesar</td><td>L06690</td><td>Diluted to 0.2 mM in E3 to prevent embryo pigmentation</td></tr><tr><td>50 mL conical tubes</td><td>Corning</td><td>352070</td><td>For heat shocking embryos</td></tr><tr><td>6 lb nylon fishing line</td><td>SecureLine</td><td>NMT250</td><td>For making embryo manipulators</td></tr><tr><td>7.5 mL transfer pipet</td><td>Globe Scientific, Inc.</td><td>135010</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma</td><td>C3881</td><td>For E3</td></tr><tr><td>Cotton swabs</td><td>Puritan</td><td>867-WC NO GLUE</td><td>For making embryo manipulators</td></tr><tr><td>Cre recombinase</td><td>New England Biolabs</td><td>M0298M</td><td></td></tr><tr><td>Digital dry bath</td><td>Genemate</td><td>490016-616</td><td>Used to store LMA at 42 &amp;deg;C</td></tr><tr><td>Epifluorescence dissection scope</td><td></td><td></td><td></td></tr><tr><td>Glass capillary tubes</td><td>World Precision Instruments</td><td>TW100F-4</td><td></td></tr><tr><td>Incubator</td><td>Forma Scientific</td><td>3158</td><td>To maintain embryos at 28 &amp;deg;C</td></tr><tr><td>Injection plate molds</td><td>Adaptive Science Tools</td><td>TU-1</td><td></td></tr><tr><td>Isotemp water bath</td><td>Fisher Scientific</td><td>2320</td><td>For heat shocking embryos</td></tr><tr><td>KCl</td><td>AMRESCO</td><td>0395</td><td>For E3 and for DNA solution for injections</td></tr><tr><td>Laser-scanning confocal microscope</td><td>Zeiss</td><td>LSM710</td><td></td></tr><tr><td>LE agarose</td><td>Genemate</td><td>E3120</td><td>To create agarose injection plates</td></tr><tr><td>Low-melt agarose (LMA)</td><td>AMRESCO</td><td>J234</td><td></td></tr><tr><td>Mating tanks</td><td>Aquaneering, Inc.</td><td>ZHCT100</td><td></td></tr><tr><td>Methylene blue</td><td>Sigma</td><td>M9140</td><td>For E3</td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>9397</td><td>For E3</td></tr><tr><td>Micromanipulator</td><td>World Precision Instruments</td><td>M3301</td><td></td></tr><tr><td>Micropipette Puller</td><td>Sutter Instrument Co.</td><td>P-97</td><td></td></tr><tr><td>MS-222 Tricaine-S</td><td>Western Chemical, Inc.</td><td></td><td>Stock made at 4 mg/mL in reverse osmosis (RO) water, then added dropwise to E3 to final concentration of 0.2 mM to anesthetize embryos</td></tr><tr><td>NaCl</td><td>J.T. Baker</td><td>4058-01</td><td>For E3</td></tr><tr><td>Petri dishes (90 mm, 60 mm)</td><td>Genesee Scientific</td><td>32-107G</td><td>To house embryos and create imaging chamber (60 mm)</td></tr><tr><td>Phenol red</td><td>Sigma</td><td>P0290</td><td></td></tr><tr><td>Soft stitch ring markers</td><td>Clover Needlecraft, Inc.</td><td>354</td><td>For creating imaging chamber with Petri dish</td></tr><tr><td>Super glue (Ultra gel control)</td><td>Loctite</td><td>1363589</td><td>For making embryo manipulators</td></tr><tr><td>Syringe needles</td><td>Beckton Dickinson</td><td>BD329412</td><td>For dechorionating embryos</td></tr></tbody></table>

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.

This section illustrates examples of results that can be obtained using the in vivo multicolor time-lapse imaging approach described here. We show that Brainbow color-coded clones of cells in the proliferative ventricular zone of the developing zebrafish hindbrain14 (Figure 1).
Typically, when Brainbow-labeled cells were arranged along a particular radial fiber, they shared the same color (Figure 1D), which cou...

Watch this Scientific Journal Video about Visualizing the Developing Brain in Living Zebrafish using Brainbow and Time-lapse Confocal Imaging at JoVE.com

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

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 ...

visualizing-developing-brain-living-zebrafish-using-brainbow-time

Research

JoVE Journal

Developmental Biology

8.5K Views.  Lewis & Clark College. 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.

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

Labeling and Imaging Cells in the Zebrafish Hindbrain

Key to understanding the morphogenetic processes that shape the early vertebrate embryo is the ability to image cells at high resolution. In zebrafish embryos, injection of plasmid DNA results in mosaic expression, allowing for the visualization of single cells or small clusters of cells 1 . We describe how injection of plasmid DNA encoding membrane-targeted Green Fluorescent Protein (mGFP) under the control of a ubiquitous promoter can be used for imaging cells undergoing neurulation. Central to this protocol is the methodology for imaging labeled cells at high resolution in sections and also in real time. This protocol entails the injection of mGFP DNA into young zebrafish embryos. Embryos are then processed for vibratome sectioning, antibody labeling and imaging with a confocal microscope. Alternatively, live embryos expressing mGFP can be imaged using time-lapse confocal microscopy. We have previously used this straightforward approach to analyze the cellular behaviors that drive neural tube formation in the hindbrain region of zebrafish embryos 2. The fixed preparations allowed for unprecedented visualization of cell shapes and organization in the neural tube while live imaging complemented this approach enabling a better understanding of the cellular dynamics that take place during neurulation.

Key to understanding the morphogenetic processes that shape the early embryo is the ability to image cells at high resolution. We describe here a technique for labeling single cells or small clusters of cells in whole zebrafish embryos with membrane-targeted Green Fluorescent Protein.

Key to understanding the morphogenetic processes that shape the early vertebrate embryo is the ability to image cells at high resolution. In zebrafish ...

Neuroscience

Time-lapse Live Imaging of Clonally Related Neural Progenitor Cells in the Developing Zebrafish Forebrain

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To understand the behavior of neural progenitor cells in the complex in vivo environment, time-lapse live imaging of neural progenitor cells in an intact brain is critically required. In this video, we exploit the unique features of zebrafish embryos to visualize the development of forebrain neural progenitor cells in vivo. We use electroporation to genetically and sparsely label individual neural progenitor cells. Briefly, DNA constructs coding for fluorescent markers were injected into the forebrain ventricle of 22 hours post fertilization (hpf) zebrafish embryos and electric pulses were delivered immediately. Six hours later, the electroporated zebrafish embryos were mounted with low melting point agarose in glass bottom culture dishes. Fluorescently labeled neural progenitor cells were then imaged for 36hours with fixed intervals under a confocal microscope using water dipping objective lens. The present method provides a way to gain insights into the in vivo development of forebrain neural progenitor cells and can be applied to other parts of the central nervous system of the zebrafish embryo.

The present video demonstrates a method which takes advantage of the combination of electroporation and confocal microscopy to perform live imaging on individual neural progenitor cells in the developing zebrafish forebrain. In vivo analysis of the development of forebrain neural progenitor cells at a clonal level can be achieved in this way.

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To ...

A Versatile Mounting Method for Long Term Imaging of Zebrafish Development

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. In particular, posterior body elongation is an essential process in embryonic development by which multiple tissue deformations act together to direct the formation of a large part of the body axis. In order to observe this process by long-term time-lapse imaging it is necessary to utilize a mounting technique that allows sufficient support to maintain samples in the correct orientation during transfer to the microscope and acquisition. In addition, the mounting must also provide sufficient freedom of movement for the outgrowth of the posterior body region without affecting its normal development. Finally, there must be a certain degree in versatility of the mounting method to allow imaging on diverse imaging set-ups. Here, we present a mounting technique for imaging the development of posterior body elongation in the zebrafish D. rerio. This technique involves mounting embryos such that the head and yolk sac regions are almost entirely included in agarose, while leaving out the posterior body region to elongate and develop normally. We will show how this can be adapted for upright, inverted and vertical light-sheet microscopy set-ups. While this protocol focuses on mounting embryos for imaging for the posterior body, it could easily be adapted for the live imaging of multiple aspects of zebrafish development.

Here, we present a versatile mounting method that allows for the long-term time-lapse imaging of the posterior body development of live zebrafish embryos without perturbing normal development.

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. ...

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 ...

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

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 ...

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 ...

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 ...