Funding for this work was supported primarily by start-up funds from Reed College to KLC, Helen Stafford Research Fellowship funds to OLH, and a Reed College Science Research Fellowship to YK. This project began in Steve Wilson&#39;s lab as a collaboration with HR, who was supported by a Wellcome Trust Studentship (2009-2014). We thank M&#225;t&#233; Varga, Steve Wilson, and other members of the Wilson lab for initial discussions about this project, and we especially thank Florencia Cavodeassi and Kate Edwards, who were the first to teach KLC how to mount embryos in agarose and perform zebrafish brain dissections. We also thank Greta Glover and Jay Ewing for help with assembling our tungsten needle-sharpening device.

The methods in this paper were conducted in accordance with guidelines and approval of the Institutional Animal Care and Use Committees of Reed College and University College London. See the Table of Materials for details about zebrafish strains used in this study.
1. Prepare materials and tools
<ol>
	<li>Make solutions.
	<ol>
		<li>Make embryo medium (E314) by diluting a 60x stock (300 mM NaCl, 10.2 mM KCl, 20 mM CaCl2-dihydrate, and 20 mM MgCl2-hexahydrate in deionized water, autoclaved, and stored at room temperature). Add 160 mL of 60x E3 to 10 L of deionized water. Store at room temperature.</li>
		<li>Make 1% low melting point (LMP) agarose in E3. Dissolve 1 g of LMP agarose into 100 mL of E3 by boiling in the microwave for 1-2 min on high power. Aliquot molten agarose into 1.7 mL microfuge tubes, and store at 4 &#176;C. Boil in a water bath and hold in a 40 &#176;C heat block when ready to use for embedding.</li>
		<li>Make Marc&#39;s Modified Ringers (MMR15) isotonic solution by diluting a 10x stock (1 M NaCl, 20 mM KCl, 10 mM MgSO4, 20 mM CaCl2-dihydrate, 50 mM HEPES, pH adjusted to 7.5 with 10 M NaOH, then autoclaved, and stored at room temperature). Add 10 mL of 10x MMR to 90 mL of deionized water.</li>
	</ol>
	</li>
	<li>Pour Sylgard plates according to the manufacturer&#39;s instructions, allowing them to set and dry for several days16.</li>
	<li>Electrolytically sharpen tungsten needles (protocol modified from 17).
	<ol>
		<li>First, prepare a 10% (w/v) KOH solution by adding 200 mL of deionized water to a wide-mouthed glass jar with a screw-top lid. Slowly stir in 20 g of KOH pellets. 
		NOTE: This caustic solution is highly exothermic. Wear gloves and eye protection, and stir the solution in the hood.</li>
		<li>Cut a 2-3 cm length of tungsten wire and insert it into the needle holder.</li>
		<li>Connect the cathode and anode wires to the power supply. Attach the alligator clip at the end of the cathode wire to a partially straightened paper clip and insert the paper clip into the KOH solution, attaching it to the side of the jar.</li>
		<li>Next, attach the alligator clip of the anode&#160;wire to the neck of the needle holder. Turn the power on (to ~20 V) and dip the tungsten wire into the KOH solution, pulling the wire out of the solution at an angle to electrolytically sharpen the wire into a fine tip. 
		NOTE: Bubbles will come from the paper clip as the reaction proceeds.</li>
		<li>Check the tip of the needle under the dissecting microscope to ensure that it is sharp enough.</li>
		<li>Rinse the needle with deionized water to remove all KOH residue. Make several needles ahead of time and keep them until the larval surgeries and dissections.</li>
	</ol>
	</li>
	<li>Make chambered slides for imaging zebrafish specimens with an upright confocal microscope.
	<ol>
		<li>Obtain glass microscope slides and two-part epoxy resin (see the Table of Materials). Mix the epoxy according to the instructions on the package, and paint thick rings or rectangles on the glass slides. Allow the slides to cure and dry in the hood for at least 24 h before use.</li>
	</ol>
	</li>
</ol>
2. Embryo collection and rearing
<ol>
	<li>Pair adult male and female fish of the desired genotypes in breeding boxes in the late afternoon/early evening. The next morning, after they have spawned, collect newly fertilized eggs with a mesh strainer and transfer them to E3 in 100 mm Petri dishes.</li>
	<li>Incubate the embryos at ~27.5 &#176;C with a 14 h light/10 h dark cycle until the day of the surgery. Change E3 daily or as needed.</li>
	<li>Feed the larvae using a dropper full of harvested rotifers once a day, beginning either at 5 days post fertilization (dpf) or one day after surgery. After the larvae have several hours to eat, change the E3 to remove any remaining rotifers and waste products. 
	&#8203;NOTE: Do not feed larvae on the day of surgery.</li>
</ol>
3. Prepare larvae for surgery
<ol>
	<li>On the day of surgery, use a wide-bore, glass Pasteur pipette to transfer 10-15 larvae to a 35 mm Petri dish filled with fresh E3.</li>
	<li>Anesthetize the larvae by adding 3-5 drops of 0.4% w/v Tricaine solution (0.4 g Tricaine dissolved in 210 mM Tris, pH 9; final solution adjusted to a pH of 7.4 if necessary18) to the dish for a final concentration of ~0.0015% w/v Tricaine. Look for lack of response to touch to determine if the larvae are adequately anesthetized. If they are still responsive to touch after ~3 min, add 2-3 more drops of 0.4% w/v Tricaine solution and reassess. 
	NOTE: At this stage of development, it is possible to monitor blood flow and heart rate in addition to motility under the dissecting microscope.</li>
	<li>Immobilize the anesthetized zebrafish larvae by embedding them in 1% LMP agarose dissolved in E3.
	<ol>
		<li>First, place the lid of a 35 mm Petri dish face up under the dissecting microscope. Next, take one larva up into a narrow-bore, glass Pasteur pipette with only a small amount of E3.</li>
		<li>Then, aspirate ~200 &#181;L of melted, warm (~40 &#176;C) 1% LMP agarose into the pipette with the larva. Finally, squirt the larva and agarose onto the upside-down Petri dish lid.</li>
		<li>Use a dull tungsten needle to quickly but gently maneuver the larva so that it is lateral, with one eye facing up. Wait for the agarose to set (~5 min). 
		&#8203;NOTE: If the agarose hardens while the larva is not lateral, free it from the agarose (as described in step 5) and return it to the dish with E3 and tricaine.</li>
	</ol>
	</li>
</ol>
4. Eye-removal
<ol>
	<li>Once the agarose is set, and the larva is positioned laterally, use the tip of a very fine, electrolytically sharpened tungsten needle to pierce the skin around the eye, following the edge of the eye orbit.</li>
	<li>Next, slide the edge of the needle (not the tip) under the eye from the temporal-ventral side of the eye. Use controlled pressure to release the eye from the socket.</li>
	<li>Use very fine surgical forceps to remove the eye by pinching the optic nerve and pushing the eye from medial to lateral. Alternatively, simply keep pressing the eye dorsally and anteriorly with the side of the needle, eventually slicing through the optic nerve and releasing the eye. 
	&#8203;NOTE: If tissues other than the eye are accidentally stabbed during the surgery, quickly and humanely kill the larva by rapid decapitation.</li>
</ol>
5. Postoperative care until experimental endpoint
<ol>
	<li>After successful eye removal, cover the agarose with MMR solution.</li>
	<li>Liberate each larva from the agarose by gently brushing a tungsten needle around their head and then around their body while stabilizing the Petri dish lid with forceps.</li>
	<li>Transfer the one-eyed larvae to a 35 mm Petri dish that contains MMR supplemented with a 1:100 dilution of a penicillin/streptomycin cocktail. Incubate the embryos at ~27.5 &#176;C with a 14 h light/10 h dark cycle. 
	&#8203;NOTE: This isotonic solution of MMR supplemented with antibiotics initially aids larval healing and survival. Each dish can hold up to 15 recovering larvae.</li>
	<li>The next day, return the larvae to E3. Starting the afternoon following surgery, feed larvae (~6 dpf and older) using a dropper full of harvested rotifers once a day. After the larvae have had several hours to eat, change the E3 to remove any remaining rotifers and waste products.</li>
</ol>
6. Fixing larvae
<ol>
	<li>Once the larvae have reached the desired developmental stage for analysis, anesthetize them with a lethal dose of tricaine (~0.4% final concentration) until their hearts have stopped.</li>
	<li>Pipette up to 30 larvae per 1.5 mL microfuge tube. Remove excess E3 and then add 0.5-1 mL of fixative solution (4% paraformaldehyde (PFA) and 4% sucrose in phosphate-buffered saline (PBS)) to fix the tissue. Incubate the larvae overnight at 4 &#176;C.</li>
	<li>Wash the larvae with PBS the next day by removing the fixative and rinsing the larvae 3-4 times with PBS. Store the larvae at 4 &#176;C for up to 7 days before dissection.</li>
</ol>
7. Dissecting larvae to reveal brains (adapted from 15)
<ol>
	<li>Suspend the fixed larvae in PBS droplets on a Sylgard plate under a dissecting microscope. Secure them laterally by placing two tungsten pins (typically old tungsten needles that are less than 2 cm) through the notochord, with one pin just posterior to the pigmented area covering the aorta-gonad-mesonephros (AGM) region and another in line with the end of the yolk extension. 
	NOTE: Position the tungsten pins in as few attempts as possible because the tissue will become more friable with each puncture.</li>
	<li>Use a very sharp tungsten needle and needle-sharp forceps to expose the brain by removing, in this order, the eye(s), ear, jaw, digestive tract, and dorsal cranial skin.
	<ol>
		<li>First, remove the eye using a technique similar to the surgical eye-removal in step 4. Then, unpin the larva and flip it to the opposite side, so the other eye is accessible and repeat. Alternatively, poke the needle through the jaw to the other side and remove the eye without unpinning. 
		NOTE: Eyes may be collected at this point if analysis of this tissue is desired (similar to 19).</li>
		<li>Next, use the tungsten needle to scratch from the temporal to the ventral side of the ear. In the same action/motion, bring the needle posterior to the jaw and gently pull anteriorly until the ear and jaw are removed.</li>
		<li>Use forceps to pull the ventral organs and any remaining yolk out.</li>
		<li>Finally, make a shallow incision in the dorsal cranial skin near the junction between the hindbrain and spinal cord. Lift the skin with the forceps and pull it anteriorly and around the telencephalon. If this proves too difficult, start at the initial incision in the hindbrain and pull the skin first laterally and then ventrally and anteriorly, taking great care not to scratch the lateral edges of the tectum. 
		NOTE: Because the midbrain is the object of study, the hindbrain can be cut; however, a deep incision may result in decapitation.</li>
		<li>Remove any remaining tissue with forceps. Unpin the larva and transfer to PBS in a 1.5 mL microfuge tube. 
		&#8203;NOTE: Leave the tail attached to the brain for better visibility when performing wholemount protocols.</li>
	</ol>
	</li>
</ol>
8. Brain dehydration and storage
<ol>
	<li>Remove PBS from the brains and add 1 mL of PBS containing 0.1% TritonX-100 (PBST). Incubate for 5-10 min at room temperature.</li>
	<li>Remove the PBST and replace it with a 50:50 methanol:PBST solution. Incubate for 5-10 min at room temperature.</li>
	<li>Wash the brains with 100% methanol (MeOH) twice for 5 min each at room temperature.</li>
	<li>Store the brains in 100% MeOH at -20 &#176;C for at least 16 h before performing immunohistochemistry or other wholemount procedures. 
	&#8203;NOTE: Brains can be stored in MeOH for up to 2 years at -20 &#176;C.</li>
</ol>
9. Immunohistochemistry
NOTE: Established protocols for many useful wholemount techniques in zebrafish can be found on ZFIN20. This manuscript provides examples comparing one-eyed and two-eyed larvae that were immunostained with antibodies that recognize either the red fluorescent protein (RFP), which is expressed in optic nerve axons in the Tg[atoh7:RFP] line (Figure 2), or phosphorylated histone H3 (PH3), which highlights mitotic cells (Figure 3). A standard immunohistochemistry protocol for wholemount embryos and larvae is summarized below.
<ol>
	<li>First, rehydrate the larval brains with a graded series of MeOH:PBST washes at room temperature by incubating the specimens in 50:50 MeOH:PBST for 5 min, then in 30:70 MeOH:PBST for 5 min, and ending with 3 washes of PBST.</li>
	<li>To permeabilize the brains, incubate them in PBST supplemented with proteinase K (PK) at a final concentration of 20 &#181;g/ml PK for 30 min at 37 &#176;C. Store aliquots of 10 mg/mL PK at -20 &#176;C and thaw them before use.</li>
	<li>Remove the PK solution and wash away any remaining enzyme by rinsing the brains with PBST 3 times over 15 min.</li>
	<li>Refix the permeabilized brains by incubating them in 4% PFA for 20 min at 25 &#176;C (or room temperature). Then, remove PFA and wash away any remaining fixative by rinsing the brains 3 times over 15 min with PBST.</li>
	<li>Replace the final PBST rinse with freshly made immunoblocking (IB) buffer (10% normal goat serum and 1% DMSO in PBST) and incubate at room temperature for at least 1 h.</li>
	<li>Dilute primary antibodies into IB buffer at the appropriate concentration (e.g., 1:500 for RFP and 1:300 for PH3 antibodies).</li>
	<li>Remove IB buffer from brains and replace it with the primary antibody dilution. Incubate overnight at 4 &#176;C on an orbital shaker with gentle rocking. Ensure tubes are securely resting on their sides.</li>
	<li>The next morning, remove the primary antibody solution with a micropipette, rinse a couple of times in PBST, and then wash the brains in PBST at room temperature for 4 x 30 min. 
	NOTE: Primary antibody dilutions can be reused once within ~7 days if stored at 4 &#176;C.</li>
	<li>Rinse the brains in IB buffer while diluting secondary antibodies into IB buffer at the appropriate concentration (e.g., 1:500 for Alexa-fluor 568 anti-rabbit).</li>
	<li>Remove the IB buffer and replace it with the secondary antibody dilution. Incubate overnight at 4 &#176;C on an orbital shaker with gentle rocking of the tubes resting on their sides.</li>
	<li>The next morning, remove the secondary antibody solution, rinse with PBST, and then wash brains in PBST at room temperature for 4 x 30 min.</li>
	<li>Counterstain with 4&#39;,6-diamidino-2-phenylindole (DAPI) or ToPro3 (1:5,000 in PBST at 4 &#176;C overnight).</li>
	<li>The next morning, transfer the larval brains into PBS and proceed to the mounting and imaging steps listed below.</li>
</ol>
10. Mounting and imaging
<ol>
	<li>Secure a chambered slide into the lid of a 100 mm Petri dish with vacuum grease.</li>
	<li>Transfer the immunostained and processed larvae to a well plate or depression slide to view them with a dissecting microscope.</li>
	<li>Mount the larvae on the chambered slide in columns of 1% LMP agarose.
	<ol>
		<li>Using a glass Pasteur pipette, put one larva on the chambered slide, depositing as little PBS as possible. With the same pipette, cover the larva with molten 1% LMP agarose (~40 &#176;C).</li>
		<li>With a blunt tungsten needle or forceps, draw the agarose into a column and then position the larva as symmetrically as possible, with the dorsal surface visible.</li>
		<li>Repeat this procedure for the remaining larvae.</li>
		<li>Once the agarose has gelled, cover it with a few drops of PBS and then place it on the stage of the confocal microscope.</li>
	</ol>
	</li>
	<li>Align the objective lens with the sample, focus the microscope using transmitted light, and illuminate the sample with the appropriate lasers. 
	NOTE: In this example, 405 nm and 568 nm wavelengths were used to excite DAPI and RFP, respectively.</li>
	<li>Adjust the gain, offset, and laser power by moving the slider bars to an appropriate level relative to the signal and the used detector. Check the histograms for each channel and click the State of saturation indicator button above the image to gauge the level of exposure, ensuring that none of the pixels are saturated and that the signal-to-noise ratio is high. For an example of optimum confocal imaging parameters for cellular and subcellular resolution within zebrafish, see 21.</li>
	<li>Set the upper and lower limits of the stack of z-planes by using the scrollwheel on the mouse to find the top and bottom of the sample. Click the top and bottom limits for image acquisition and then trigger the z-stack capture by clicking the Run now button. 
	NOTE: Depending on how symmetrically the brain is mounted, the total z-depth for a 9 dpf larval zebrafish brain will be ~150-200 &#181;m.</li>
	<li>Process and colorize the images with colorblind accessibility in mind (e.g., use blue and orange or magenta and green for two-color images). Set lookup tables in the software used to capture the images or image processing software such as Photoshop from Adobe.</li>
	<li>In Photoshop, color raw photomicrographs saved as 8 bit Tagged Image File Format (TIFF) by (i) converting the Image Mode to RGB and (ii) accessing the Curves option under the Image | Adjustments menus and then (iii) sliding the appropriate colored light outputs to 0 or 255 levels to achieve the desired color. For example, to achieve a green signal, select the Red channel and slide its output to 0; then, select the Blue channel and slide its output to 0 too. Similarly, to achieve a magenta signal, select the Green channel and slide its output to 0; leave the red and blue channels as they are. 
	NOTE: Similar steps can be performed in the free Gnu Image Manipulation Program (GIMP).</li>
	<li>Manually quantify the numbers of cells displaying a specific marker using the cell counter plugin in ImageJ22, available under Analyze in the Plugins menu. Examples of how to use ImageJ for cell counting can be found in many tutorials, including 23.</li>
	<li>Generate graphical representations of data in statistical software such as RStudio (see supplemental data for .Rmd file).</li>
</ol>

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Fiji Available from: <a target="_blank" href="https://figi.sc/">https://figi.sc/</a> (2021)</li><li>O'Brien, J., Hayder, H., Peng, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Automated+quantification+and+analysis+of+cell+counting+procedures+using+ImageJ+plugins.">Automated quantification and analysis of cell counting procedures using ImageJ plugins.</a> Journal of Visualized Experiments: JoVE. (117), e54719 (2016).</li><li>Poggi, L., Vitorino, M., Masai, I., Harris, W. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Influences+on+neural+lineage+and+mode+of+division+in+the+zebrafish+retina+in+vivo.">Influences on neural lineage and mode of division in the zebrafish retina in vivo.</a> The Journal of Cell Biology. 171 (6), 991-999 (2005).</li><li>Karlstrom, R. O., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+mutations+affecting+retinotectal+axon+pathfinding.">Zebrafish mutations affecting retinotectal axon pathfinding.</a> Development. 123 (1), 427-438 (1996).</li><li>Harvey, B. M., Baxter, M., Granato, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optic+nerve+regeneration+in+larval+zebrafish+exhibits+spontaneous+capacity+for+retinotopic+but+not+tectum+specific+axon+targeting.">Optic nerve regeneration in larval zebrafish exhibits spontaneous capacity for retinotopic but not tectum specific axon targeting.</a> PLOS ONE. 14 (6), 0218667 (2019).</li><li>Robles, E., Filosa, A., Baier, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Precise+lamination+of+retinal+axons+generates+multiple+parallel+input+pathways+in+the+tectum.">Precise lamination of retinal axons generates multiple parallel input pathways in the tectum.</a> Journal of Neuroscience. 33 (11), 5027-5039 (2013).</li><li>Vargas, M. E., Barres, B. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Why+Is+Wallerian+degeneration+in+the+CNS+so+slow.">Why Is Wallerian degeneration in the CNS so slow.</a> Annual Review of Neuroscience. 30 (1), 153-179 (2007).</li><li>Hughes, A. N., Appel, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microglia+phagocytose+myelin+sheaths+to+modify+developmental+myelination.">Microglia phagocytose myelin sheaths to modify developmental myelination.</a> Nature Neuroscience. 23 (9), 1055-1066 (2020).</li><li>de Calbiac, H., Dabacan, A., Muresan, R., Kabashi, E., Ciura, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Behavioral+and+physiological+analysis+in+a+zebrafish+model+of+epilepsy.">Behavioral and physiological analysis in a zebrafish model of epilepsy.</a> Journal of Visualized Experiments: JoVE. (176), e58837 (2021).</li><li>Adams, S. L., Zhang, T., Rawson, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effect+of+external+medium+composition+on+membrane+water+permeability+of+zebrafish+(Danio+rerio)+embryos.">The effect of external medium composition on membrane water permeability of zebrafish (Danio rerio) embryos.</a> Theriogenology. 64 (7), 1591-1602 (2005).</li><li>Fredj, N. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptic+activity+and+activity-dependent+competition+regulates+axon+arbor+maturation,+growth+arrest,+and+territory+in+the+retinotectal+projection.">Synaptic activity and activity-dependent competition regulates axon arbor maturation, growth arrest, and territory in the retinotectal projection.</a> Journal of Neuroscience. 30 (32), 10939-10951 (2010).</li><li>Alberio, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+light-gated+potassium+channel+for+sustained+neuronal+inhibition.">A light-gated potassium channel for sustained neuronal inhibition.</a> Nature Methods. 15 (11), 969-976 (2018).</li><li>Kay, J. N., Finger-Baier, K. C., Roeser, T., Staub, W., Baier, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Retinal+ganglion+cell+genesis+requires+lakritz,+a+zebrafish+atonal+homolog.">Retinal ganglion cell genesis requires lakritz, a zebrafish atonal homolog.</a> Neuron. 30 (3), 725-736 (2001).</li><li>Gnuegge, L., Schmid, S., Neuhauss, S. C. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+the+activity-deprived+zebrafish+mutant+macho+reveals+an+essential+requirement+of+neuronal+activity+for+the+development+of+a+fine-grained+visuotopic+map.">Analysis of the activity-deprived zebrafish mutant macho reveals an essential requirement of neuronal activity for the development of a fine-grained visuotopic map.</a> The Journal of Neuroscience. 21 (10), 3542-3548 (2001).</li><li>Jeffery, W. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Astyanax+surface+and+cave+fish+morphs.">Astyanax surface and cave fish morphs.</a> EvoDevo. 11 (1), 14 (2020).</li><li>Sieger, D., Peri, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+for+studying+microglia:+The+first,+the+popular,+and+the+new.">Animal models for studying microglia: The first, the popular, and the new.</a> Glia. 61 (1), 3-9 (2013).</li><li>Svahn, A. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+ramified+microglia+from+early+macrophages+in+the+zebrafish+optic+tectum.">Development of ramified microglia from early macrophages in the zebrafish optic tectum.</a> Developmental Neurobiology. 73 (1), 60-71 (2013).</li><li>Herzog, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+clearance+of+cellular+debris+by+microglia+limits+secondary+neuronal+cell+death+after+brain+injury+in+vivo.">Rapid clearance of cellular debris by microglia limits secondary neuronal cell death after brain injury in vivo.</a> Development. 146 (9), (2019).</li><li>Chen, J., Poskanzer, K. E., Freeman, M. R., Monk, K. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live-imaging+of+astrocyte+morphogenesis+and+function+in+zebrafish+neural+circuits.">Live-imaging of astrocyte morphogenesis and function in zebrafish neural circuits.</a> Nature Neuroscience. 23 (10), 1297-1306 (2020).</li></ol>

The authors have no conflicts of interest to disclose.

The techniques described in this paper illustrate one of many approaches for studying vertebrate visual system development in zebrafish. Other researchers have published methods to dissect the embryonic retina and perform gene expression analyses19 or visualize neuronal activity in the optic tectum30. This paper provides an approach for exploring how differential retinal input may influence cell behaviors in the optic tectum.
To ensure successful...

The goal of this method is to investigate how retinal input influences the growth and development of the optic tectum, the visual processing center in the zebrafish brain. By removing one eye and then comparing the two sides of the optic tectum, tectal changes within the same specimen can be observed and normalized, enabling comparison across multiple specimens. Modern molecular approaches combined with this technique will yield insights into the mechanisms underlying visual system growth and development, as well as axonal degeneration and regeneration.
Sensory systems - visual, auditory, and somatosensory - gather information from external organs and relay that information to the central nervous system, generating &#34;maps&#34; of the external world across the midbrain1,2. Vision is the dominant sensory modality for nearly all vertebrates, including many fishes. The retina, the neural tissue in the eye, gathers information with a neuronal circuit consisting primarily of photoreceptors, bipolar cells, and retinal ganglion cells (RGCs), the projection neurons of the retina. RGCs have long axons that find their way across the inner surface of the retina to the optic nerve head, where they fasciculate and travel together through the brain, ultimately terminating in the visual processing center in the dorsal midbrain. This structure is called the optic tectum in fish and other non-mammalian vertebrates and is homologous to the superior colliculus in mammals3.
The optic tectum is a bilaterally symmetric multilayered structure in the dorsal midbrain. In zebrafish and most other fishes, each lobe of the optic tectum receives visual input solely from the contralateral eye, such that the left optic nerve terminates in the right tectal lobe and the right optic nerve terminates in the left tectal lobe4 (Figure 1). Like its mammalian counterpart, the superior colliculus, the optic tectum integrates visual information with other sensory inputs, including audition and somatosensation, controlling shifts in visual attention and eye movements such as saccades1,5,6. However, unlike the mammalian superior colliculus, the optic tectum continuously generates new neurons and glia from a specialized stem cell niche near the medial and caudal edges of the tectal lobes called the tectal proliferation zone7. Maintenance of proliferative progenitors in the optic tectum and other regions of the central nervous system contributes, in part, to the remarkable regenerative capacity documented in zebrafish8.
Previous work examining the brains of blind or one-eyed fishes revealed that optic tectum size is directly proportional to the amount of retinal innervation it receives9,10,11. In adult cave fish, whose eyes degenerate in early embryogenesis, the optic tectum is noticeably smaller than that of closely related, sighted surface fish9. Cave fish eye degeneration can be blocked by replacing the endogenous lens with a lens from a surface fish during embryogenesis. When these one-eyed cave fish are reared to adulthood, the innervated tectal lobe contains approximately 10% more cells than the non-innervated tectal lobe9. Similarly, in larval killifish that were incubated with chemical treatments to generate eyes of different sizes within the same individual, the side of the tectum with more innervation was larger and contained more neurons10. Evidence from optic nerve crush experiments in adult goldfish indicates that innervation promotes proliferation, with tectal cell proliferation decreasing when innervation was disrupted11.
Confirming and extending these classical studies, several recent reports provide data suggesting that proliferation in response to innervation is modulated, at least in part, by the BDNF-TrkB pathway12,13. Many open questions about optic tectum growth and development remain, including how a developing sensory system copes with injury and axon degeneration, which cellular and molecular signals enable retinal input to regulate optic tectum growth, when these mechanisms become active, and whether innervation-linked proliferation and differentiation enable the retina and its target tissue to coordinate growth rates and ensure accurate retinotopic mapping. In addition, there are much larger questions about activity-dependent development that can be addressed by interrogating the zebrafish visual system with surgical approaches such as the one described below.
To investigate the cellular and molecular mechanisms by which neural activity, specifically from visual input, alters cell survival and proliferation, the described approach directly compares innervated and denervated tectal lobes (Figure 1) within individual zebrafish larvae. This method allows for the documentation of RGC axon degeneration in the optic tectum and confirmation that the number of mitotic cells correlates with innervation.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63509/63509fig01.jpg" /> 
Figure 1: Sketches of zebrafish larvae before and after unilateral eye removal. (A) Drawing of 5 dpf larvae as viewed under a dissecting microscope. Each larva is embedded in low-melting-point agarose and oriented laterally before a tungsten needle with a sharp, hooked tip is used to scoop out the eye facing up (left eye in this example). (B) Drawing of the dorsal view of a 9 dpf larva resulting from the surgery depicted in A. Only three highly schematized RGC axons from the right eye are shown defasciculating and connecting with neurons in the left tectal lobe. Abbreviations: dpf = days post fertilization; dps = days post surgery; RGC = retinal ganglion cells. <a href="https://www.jove.com/files/ftp_upload/63509/63509fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Equipment and supplies:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Breeding boxes</td>
			<td>Aquaneering</td>
			<td>ZHCT100</td>
			<td></td>
		</tr>
		<tr>
			<td>Dow Corning high vacuum grease</td>
			<td>Sigma or equivalent supplier</td>
			<td>Z273554</td>
			<td></td>
		</tr>
		<tr>
			<td>Erlenmeyer flasks (125 mL)</td>
			<td></td>
			<td></td>
			<td>For making Marc&#39;s Modified Ringers (MMR) with antibiotics for post-surgery incubation</td>
		</tr>
		<tr>
			<td>Fine forceps - Dumont #5</td>
			<td>Fine Science Tools (FST)</td>
			<td>11252-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Pasteur pipettes</td>
			<td>DWK Lifescience</td>
			<td>63A53 &#38; 63A53WT</td>
			<td>For pipetting embryos and larvae</td>
		</tr>
		<tr>
			<td>Glass slides for microscopy</td>
			<td>VWR or equivalent supplier</td>
			<td>48311-703</td>
			<td>Standard glass microscope slides can be ordered from many different laboratory suppliers.</td>
		</tr>
		<tr>
			<td>Glassware including graduated bottles and graduated cylinders</td>
			<td></td>
			<td></td>
			<td>For making and storing solutions</td>
		</tr>
		<tr>
			<td>2-part epoxy resin</td>
			<td>ACE Hardware or other equivalent supplier of Gorilla Glue or equivalent</td>
			<td>0.85 oz syringe</td>
			<td>https://www.acehardware.com/departments/paint-and-supplies/tape-glues-and-adhesives/glues-and-epoxy/1590793</td>
		</tr>
		<tr>
			<td>Microcentrifuge tube (1.7 mL)</td>
			<td>VWR or equivalent supplier</td>
			<td>22234-046</td>
			<td></td>
		</tr>
		<tr>
			<td>Nickel plated pin holder (17 cm length)</td>
			<td>Fine Science Tools (FST)</td>
			<td>26018-17</td>
			<td>To hold tungsten wire while sharpening and performing surgeries/dissections.</td>
		</tr>
		<tr>
			<td>Nylon mesh tea strainer or equivalent</td>
			<td>Ali Express or equivalent</td>
			<td></td>
			<td>For harvesting zebrafish eggs after spawning; https://www.aliexpress.com/item/1005002219569756.html</td>
		</tr>
		<tr>
			<td>Paper clip</td>
			<td></td>
			<td></td>
			<td>For Tungsten needle sharpening device.</td>
		</tr>
		<tr>
			<td>Petri dishes 100 mm</td>
			<td>Fischer Scientific or equivalent supplier</td>
			<td>50-190-0267</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dishes 35 mm</td>
			<td>Fischer Scientific or equivalent supplier</td>
			<td>08-757-100A</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette pump</td>
			<td>SP Bel-Art or equivalent</td>
			<td>F37898-0000</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium hydroxide (KOH)</td>
			<td>Sigma</td>
			<td>909122</td>
			<td>For Tungsten needle sharpening device. Make a 10% w/v solution of KOH in the hood by adding pellets to deionized water.</td>
		</tr>
		<tr>
			<td>Power supply (variable voltage)</td>
			<td></td>
			<td></td>
			<td>For Tungsten needle sharpening device. Any power supply with variable voltage will work (even one used for gel electrophoresis).</td>
		</tr>
		<tr>
			<td>Sylgard 184 Elastomer kit</td>
			<td>Dow Corning</td>
			<td>3097358</td>
			<td></td>
		</tr>
		<tr>
			<td>Tungsten wire (0.125 mm diameter)</td>
			<td>World Precision Instruments (WPI)</td>
			<td>TGW0515</td>
			<td>Sharpen to remove eye and dissect larvae.</td>
		</tr>
		<tr>
			<td>Variable temperature heat block</td>
			<td>The Lab Depot or equivalent supplier</td>
			<td>BSH1001 or BSH1002</td>
			<td>Set to 40-42 &#176;C ahead of experiments.</td>
		</tr>
		<tr>
			<td>Wide-mouth glass jar with lid (e.g., clean jam or salsa jar)</td>
			<td></td>
			<td></td>
			<td>For Tungsten needle sharpening device.</td>
		</tr>
		<tr>
			<td>Wires with alligator clip leads</td>
			<td></td>
			<td></td>
			<td>For Tungsten needle sharpening device.</td>
		</tr>
		<tr>
			<td>Microscopes:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting microscope</td>
			<td></td>
			<td></td>
			<td>Any type will work but having adjustable transmitted light on a mirrored base is preferred.</td>
		</tr>
		<tr>
			<td>Laser scanning confocal microscope</td>
			<td></td>
			<td></td>
			<td>High NA, 20-25x water dipping objective lens is recommended. 
			Microscope control and image capture software (Elements) is used here but any confocal microscope will work.</td>
		</tr>
		<tr>
			<td>Reagents for surgeries and dissections:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate</td>
			<td>Sigma</td>
			<td>C7902</td>
			<td>For Marc&#39;s Modified Ringers (MMR) and embryo medium (E3).</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>H7006</td>
			<td>For Marc&#39;s Modified Ringers (MMR).</td>
		</tr>
		<tr>
			<td>Low melting point agarose</td>
			<td>Invitrogen</td>
			<td>16520-050</td>
			<td>Make 1% in embryo medium (E3) or Marc&#39;s Modified Ringers (MMR).</td>
		</tr>
		<tr>
			<td>Magnesium chloride hexahydrate</td>
			<td>Sigma</td>
			<td>1374248</td>
			<td>For embryo medium (E3).</td>
		</tr>
		<tr>
			<td>Magnesium sulfate</td>
			<td>Sigma</td>
			<td>M7506</td>
			<td>For Marc&#39;s Modified Ringers (MMR).</td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>19210</td>
			<td>Dilute 8% (w/v) stock with 2x concentrated PBS (diluted from 10x PBS stock).</td>
		</tr>
		<tr>
			<td>Penicillin/Streptomycin</td>
			<td>Sigma</td>
			<td>P4333-20ML</td>
			<td>Dilute 1:100 in Marc&#39;s Modified Ringers.</td>
		</tr>
		<tr>
			<td>Phosphate buffered saline (PBS) tablets</td>
			<td>Diagnostic BioSystems</td>
			<td>DMR E404-01</td>
			<td>Make 10x stock in deionized water, autoclave and store at room temperature. Dilute to 1x working concentration.</td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Sigma</td>
			<td>P3911</td>
			<td>For Marc&#39;s Modified Ringers (MMR) and embryo medium (E3).</td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Sigma</td>
			<td>S9888</td>
			<td>For Marc&#39;s Modified Ringers (MMR) and embryo medium (E3).</td>
		</tr>
		<tr>
			<td>Sodium hydroxide</td>
			<td>Sigma</td>
			<td>S5881</td>
			<td>Make 10 M and use to adjust pH of MMR to 7.4.</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma</td>
			<td>S9378</td>
			<td></td>
		</tr>
		<tr>
			<td>Tricaine-S</td>
			<td>Pentair</td>
			<td>100G #TRS1</td>
			<td>Recipe: https://zfin.atlassian.net/wiki/spaces/prot/pages/362220023/TRICAINE</td>
		</tr>
		<tr>
			<td>Reagents for immunohistochemistry:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Alexafluor 568 tagged Secondary antibody to detect rabbit IgG</td>
			<td>Invitrogen</td>
			<td>A-11011</td>
			<td>Use at 1:500 dilution for wholemount immunohistochemistry.</td>
		</tr>
		<tr>
			<td>DAPI or ToPro3</td>
			<td>Invitrogen</td>
			<td>1306 or T3605</td>
			<td>Make up 1 mg/mL solutions in DMSO; 1:5,000 dilution for counterstaining.</td>
		</tr>
		<tr>
			<td>Dimethyl sulfoxide (DMSO)</td>
			<td>Sigma</td>
			<td>D8418</td>
			<td>A component of immunoblock buffer.</td>
		</tr>
		<tr>
			<td>Methanol (MeOH)</td>
			<td>Sigma</td>
			<td>34860</td>
			<td>Mixing MeOH with aqueous solutions like PBST is exothermic. Make the MeOH/PBST solutions at least several hours ahead of time or cool them on ice before using.</td>
		</tr>
		<tr>
			<td>Normal goat serum</td>
			<td>ThermoFisher Scientific</td>
			<td>50-062Z</td>
			<td>A component of immunoblock buffer. Can be aliquoted in 1-10 mL volumes and stored at -20 &#176;C.</td>
		</tr>
		<tr>
			<td>Primary antibody to detect phosphohistone H3</td>
			<td>Millipore</td>
			<td>06-570</td>
			<td>Use at 1:300 dilution for wholemount immunohistochemistry.</td>
		</tr>
		<tr>
			<td>Primary antibody to detect Red Fluorescent Protein (RFP; detects dsRed derivatives)</td>
			<td>MBL International</td>
			<td>PM005</td>
			<td>Use at 1:500 dilution for wholemount immunohistochemistry.</td>
		</tr>
		<tr>
			<td>Proteinase K (PK)</td>
			<td>Sigma</td>
			<td>P2308-10MG</td>
			<td>Make up 10 mg/mL stock solutions in PBS and use at 10 &#181;g/mL.</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma</td>
			<td>T8787</td>
			<td>Useful to make a 20% (v/v) stock solution in PBS.</td>
		</tr>
		<tr>
			<td>Software for data analysis</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ (Fiji)</td>
			<td></td>
			<td></td>
			<td>freeware for image analysis; https://imagej.net/software/fiji/</td>
		</tr>
		<tr>
			<td>Rstudio</td>
			<td></td>
			<td></td>
			<td>freeware for statistical analysis and data visualization; https://www.rstudio.com/products/rstudio/download/</td>
		</tr>
		<tr>
			<td>Adobe Photoshop or GIMP</td>
			<td></td>
			<td></td>
			<td>Proprietary image processing software (Adobe Photoshop and Illustrator) are often used to compose figures). A freeware alternative is Gnu Image Manipulation Program (GIMP; https://www.gimp.org/)</td>
		</tr>
		<tr>
			<td>Zebrafish strains</td>
			<td></td>
			<td></td>
			<td>available from the&#160; Zebrafish International Resource Centers in the US (https://zebrafish.org/home/guide.php) or in Europe (https://www.ezrc.kit.edu/). Specialized transgenic strains that have not yet been deposited in either resource center can be requested from individual labs after publication.</td>
		</tr>
	</tbody>
</table>

eye removal in living zebrafish larvae to examine innervation-dependent growth and development of the visual system

Zebrafish exhibit remarkable life-long growth and regenerative abilities. For example, specialized stem cell niches established during embryogenesis support continuous growth of the entire visual system, both in the eye and the brain. Coordinated growth between the retinae and the optic tectum ensures accurate retinotopic mapping as new neurons are added in the eyes and brain. To address whether retinal axons provide crucial information for regulating tectal stem and progenitor cell behaviors such as survival, proliferation, and/or differentiation, it is necessary to be able to compare innervated and denervated tectal lobes within the same animal and across animals. 
Surgical removal of one eye from living larval zebrafish followed by observation of the optic tectum achieves this goal. The accompanying video demonstrates how to anesthetize larvae, electrolytically sharpen tungsten needles, and use them to remove one eye. It next shows how to dissect brains from fixed zebrafish larvae. Finally, the video provides an overview of the protocol for immunohistochemistry and a demonstration of how to mount stained embryos in low-melting-point agarose for microscopy.

To confirm whether eye removal was complete and assess how the optic tectum changes, surgeries were performed in the Tg[atoh7:RFP] strain, which labels all RGCs with a membrane-targeted RFP and, thus, all axons that project from the retina and form the optic nerve24. Although using this strain is not absolutely necessary, it enables straightforward observation and visualization of the optic nerve termini in the optic tectum neuropil. Other approaches for labeling the optic nerve, such as ...

Watch this Scientific Journal Video about Eye Removal in Living Zebrafish Larvae to Examine Innervation-dependent Growth and Development of the Visual System at JoVE.com

Eye Removal in Living Zebrafish Larvae to Examine Innervation-dependent Growth and Development of the Visual System

The article explains how to surgically remove eyes from living zebrafish larvae as the first step toward investigating how retinal input influences optic tectum growth and development. In addition, the article provides information about larval anesthetization, fixation, and brain dissection, followed by immunohistochemistry and confocal imaging.

Zebrafish exhibit remarkable life-long growth and regenerative abilities. For example, specialized stem cell niches established during embryogenesis ...

eye-removal-living-zebrafish-larvae-to-examine-innervation-dependent

Research

JoVE Journal

Developmental Biology

3.7K Views.  Reed College. The article explains how to surgically remove eyes from living zebrafish larvae as the first step toward investigating how retinal input influences optic tectum growth and development. In addition, the article provides information about larval anesthetization, fixation, and brain dissection, followed by immunohistochemistry and confocal imaging.

Video: Eye Removal in Living Zebrafish Larvae to Examine Innervation-dependent Growth and Development of the Visual System

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion (FDAP)

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological systems. In FDAP experiments, the clearance of proteins within living organisms can be measured. A protein of interest is tagged with a photoconvertible fluorescent protein, expressed in vivo and photoconverted, and the decrease in the photoconverted signal over time is monitored. The data is then fitted with an appropriate clearance model to determine the protein half-life. Importantly, the clearance kinetics of protein populations in different compartments of the organism can be examined separately by applying compartmental masks. This approach has been used to determine the intra- and extracellular half-lives of secreted signaling proteins during zebrafish development. Here, we describe a protocol for FDAP experiments in zebrafish embryos. It should be possible to use FDAP to determine the clearance kinetics of any taggable protein in any optically accessible organism.

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion FDAP

Protein levels in cells and tissues are often tightly regulated by the balance of protein production and clearance. Using Fluorescence Decay After Photoconversion (FDAP), the clearance kinetics of proteins can be experimentally measured in vivo.

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological ...

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

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 ...

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM compared to confocal systems are its low phototoxicity, gentle mounting strategies, fast acquisition with high signal to noise ratio and the possibility of imaging samples from various angles (views) for long periods of time. Imaging from multiple views unleashes the full potential of LSFM, but at the same time it can create terabyte-sized datasets. Processing such datasets is the biggest challenge of using LSFM. In this protocol we outline some solutions to this problem. Until recently, LSFM was mostly performed in laboratories that had the expertise to build and operate their own light sheet microscopes. However, in the last three years several commercial implementations of LSFM became available, which are multipurpose and easy to use for any developmental biologist. This article is primarily directed to those researchers, who are not LSFM technology developers, but want to employ LSFM as a tool to answer specific developmental biology questions. 
Here, we use imaging of zebrafish eye development as an example to introduce the reader to LSFM technology and we demonstrate applications of LSFM across multiple spatial and temporal scales. This article describes a complete experimental protocol starting with the mounting of zebrafish embryos for LSFM. We then outline the options for imaging using the commercially available light sheet microscope. Importantly, we also explain a pipeline for subsequent registration and fusion of multiview datasets using an open source solution implemented as a Fiji plugin. While this protocol focuses on imaging the developing zebrafish eye and processing data from a particular imaging setup, most of the insights and troubleshooting suggestions presented here are of general use and the protocol can be adapted to a variety of light sheet microscopy experiments.

Light sheet fluorescence microscopy is an excellent tool for imaging embryonic development. It allows recording of long time-lapse movies of live embryos in near physiological conditions. We demonstrate its application for imaging zebrafish eye development across wide spatio-temporal scales and present a pipeline for fusion and deconvolution of multiview datasets.

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM ...

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize mammalian adaptive immunity. Drosophila and mammalian hematopoiesis occur in spatially and temporally distinct phases to produce several blood cell lineages. Both systems maintain reservoirs of blood cell progenitors with which to expand or replace mature lineages. The hematopoietic system allows Drosophila and mammals to respond to and to adapt to immune challenges. Importantly, the transcriptional regulators and signaling pathways that control the generation, maintenance, and function of the hematopoietic system are conserved from flies to mammals. These similarities allow Drosophila to be used to genetically model hematopoietic development and disease.
Here we detail assays to examine the hematopoietic system of Drosophila larvae. In particular, we outline methods to measure blood cell numbers and concentration, visualize a specific mature lineage in vivo, and perform immunohistochemistry on blood cells in circulation and in the hematopoietic organ. These assays can reveal changes in gene expression and cellular processes including signaling, survival, proliferation, and differentiation and can be used to investigate a variety of questions concerning hematopoiesis. Combined with the genetic tools available in Drosophila, these assays can be used to evaluate the hematopoietic system upon defined genetic alterations. While not specifically outlined here, these assays can also be used to examine the effect of environmental alterations, such as infection or diet, on the hematopoietic system.

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Drosophila and mammalian hematopoietic systems share many common features, making Drosophila an attractive genetic model to study hematopoiesis. Here we demonstrate dissection and mounting of the major larval hematopoietic organ for immunohistochemistry. We also describe methods to assay various larval hematopoietic compartments including circulating hemocytes and sessile crystal cells.

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize ...

Induction of Hypoxia in Living Frog and Zebrafish Embryos

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and zebrafish embryos. Our system comprises a chamber featuring a simple setup that is nevertheless robust to induce and maintain a specific oxygen concentration and temperature in any experimental solution of choice. The presented system is very cost-effective but highly functional, it allows induction and sustainment of hypoxia for direct experiments in vivo and for various time periods up to 48 h.
To monitor and study the effects of hypoxia, we have employed two methods - measurement of levels of hypoxia-inducible factor 1alpha (HIF-1&#945;) in whole embryos or specific tissues and determination of retinal stem cell proliferation by 5-ethynyl-2&#8242;-deoxyuridine (EdU) incorporation into the DNA. HIF-1&#945; levels can serve as a general hypoxia marker in the whole embryo or tissue of choice, here embryonic retina. EdU incorporation into the proliferating cells of embryonic retina is a specific output of hypoxia induction. Thus, we have shown that hypoxic embryonic retinal progenitors decrease proliferation within 1 h of incubation under 5% oxygen of both frog and zebrafish embryos.
Once mastered, our setup can be employed for use with small aquatic model organisms, for direct in vivo experiments, any given time period and under normal, hypoxic or hyperoxic oxygen concentration or under any other given gas mixture.

We introduce a novel hypoxic chamber system for use with aquatic organisms such as frog and zebrafish embryos. Our system is simple, robust, cost-effective and allows the induction and sustainment of hypoxia in vivo and for up to 48 h. We present 2 reproducible methods to monitor the effectiveness of hypoxia.

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and ...

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 ...

Single-cell Photoconversion in Living Intact Zebrafish

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause disease when disrupted. Scientists have developed clever techniques to investigate characteristics and natural dynamics of these cells within intact tissue by expressing fluorescent proteins in subsets of cells. However, at times, experiments require more selected visualization of cells within the tissue, sometimes at the single-cell or population-of-cells manner. To achieve this and visualize single cells within a population of cells, scientists have utilized single-cell photoconversion of fluorescent proteins. To demonstrate this technique, we show here how to direct UV light to an Eos-expressing cell of interest in an intact, living zebrafish. We then image those photoconverted Eos+ cells 24 h later to determine how they changed in the tissue. We describe two techniques: single cell photoconversion and photoconversions of populations of cell. These techniques can be used to visualize cell-cell interactions, cell-fate and differentiation, and cell migrations, making it a technique that is applicable in numerous biological questions.

Here, we present a protocol to show how cell photoconversion is achieved through UV exposure to specific areas expressing the fluorescent protein, Eos, in living animals.

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause ...

Whole-Mount In Situ Hybridization in Zebrafish Embryos and Tube Formation Assay in iPSC-ECs to Study the Role of Endoglin in Vascular Development

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in zebrafish during different developmental stages. To investigate the role of endoglin(eng) in vessel formation during the development of hereditary hemorrhagic telangiectasia&#160;(HHT), morpholino-mediated targeted knockdown of eng in zebrafish are used to study its temporal expression and associated functions. Here, whole-mount in situ RNA hybridization (WISH) is employed for the analysis of eng and its downstream genes in zebrafish embryos. Also, tube formation assays are performed in HHT patient-derived induced pluripotent stem cell-differentiated&#160;endothelial cells (iPSC-ECs; with eng mutations). A specific signal amplifying system using the whole amount In Situ Hybridization &#8211; WISH provides higher resolution and lower background results compared to traditional methods. To obtain a better signal, the post-fixation time is adjusted to 30 min after probe hybridization. Because fluorescence staining is not sensitive in zebrafish embryos, it is replaced with diaminobezidine (DAB) staining here. In this protocol, HHT&#160;patient-derived iPSC lines containing an eng mutation are differentiated into endothelial cells. After coating a plate with basement membrane matrix for 30 min at 37 &#176;C, iPSC-ECs are seeded as a monolayer into wells and kept at 37 &#176;C for 3 h. Then, the tube length and number of branches are calculated using microscopic images. Thus, with this improved WISH protocol, it is shown that reduced eng expression affects endothelial progenitor formation in zebrafish embryos. This is further confirmed by tube formation assays using iPSC-ECs derived from a patient with HHT. These assays confirm the role for eng in early vascular development.

Presented here is a protocol for whole-mount in situ RNA hybridization analysis in zebrafish and tube formation assay in patient-derived induced pluripotent stem cell-derived endothelial cells to study the role of endoglin in vascular formation.

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in ...

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. Although many immunohistochemistry protocols have been optimized for mammalian tissue and tissue sections, these protocols often require modification and optimization for non-mammalian model organisms. Zebrafish are increasingly used as a model system in basic, biomedical, and translational research to investigate the molecular, genetic, and cell biological mechanisms of developmental processes. Zebrafish offer many advantages as a model system but also require modified techniques for optimal protein detection. Here, we provide our protocol for whole-mount fluorescence immunohistochemistry in zebrafish embryos and larvae. This protocol additionally describes several different mounting strategies that can be employed and an overview of the advantages and disadvantages each strategy provides. We also describe modifications to this protocol to allow detection of chromogenic substrates in whole mount tissue and fluorescence detection in sectioned larval tissue. This protocol is broadly applicable to the study of many developmental stages and embryonic structures.

Here, we present a protocol for fluorescent antibody-mediated detection of proteins in whole preparations of zebrafish embryos and larvae.

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. ...

Eye Removal in Living Zebrafish Larvae to Examine Innervation-dependent Growth and Development of the Visual System

Summary

Explore More Videos

Chapters in this video

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion (FDAP)

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

Imaging Subcellular Structures in the Living Zebrafish Embryo

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Induction of Hypoxia in Living Frog and Zebrafish Embryos

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

Single-cell Photoconversion in Living Intact Zebrafish

Whole-Mount In Situ Hybridization in Zebrafish Embryos and Tube Formation Assay in iPSC-ECs to Study the Role of Endoglin in Vascular Development

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae