Name: eye removal in living zebrafish larvae to examine innervation-dependent growth and development of the visual system
Uploaded: 2022-02-11T14:00:00.000+00:00
Duration: 9 min
Description: 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

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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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 eye extirpation and larval survival after surgery, it is important that the larvae are healthy, adequately anesthetized, and immobilized in 1% agarose. It is also crucial that forceps and tungsten needles are very sharp and clean. Larvae survive best when they are not fed 24 h before or after the surgery, and rearing larvae in MMR supplemented with antibiotics speeds the healing process. Importantly, the higher ionic strength of MMR and especially the higher concentration of calcium promotes healing14. However, longer exposure to the higher salinity can decrease survival, similar to what has been documented in embryonic zebrafish31. A most critical step is fixation to ensure successful brain dissections, which are essential for data analysis of images of immunostained samples. Using freshly made 4% PFA supplemented with 4% sucrose (affectionately called sweet fix) tends to increase the ease of skin removal and brain tissue preservation. As with the surgeries, sharp and clean tools are a must for dissecting brains.
One of the biggest challenges of this protocol is embedding larvae for eye extirpations. It is important that each person find the way that works best for them so that they can confidently and quickly remove the larval eye. If the larva is accidentally stabbed during the surgery, humanely kill it by decapitation. Another challenge is determining the most effective antibody concentrations for wholemount immunostaining. When determining the appropriate antibody concentrations, use the published literature as a starting point and optimize the protocol as using published reagents on different tissues (especially on larger tissues or older animals) may require increased concentrations and/or incubation times.
The major limitation of this method is the time a single experiment takes from start to finish and the number of fish that can be studied at any given time. For example, all the data shown in Figure 3 took about one month from the initial pairing of fish to the final data collection and analysis. This relatively low-throughput approach could be complemented by heterologous genetic approaches that either selectively silence neural activity in the visual system32,33 or by mutants that lack RGC axons or activity34,35.
The significance of this method is that it brings a traditional embryological technique to bear on modern transgenic fish lines, allowing direct comparison of cellular and genetic mechanisms operating in each of the tectal lobes within the same individual. Moreover, this method is ripe for application to other experimental systems, including Astyanax surface fish36 and/or transgenic zebrafish larvae with fluorescently-labeled microglia37,38,39 and astrocytes40 to study how the developing fish nervous system responds to such a dramatic injury.

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.remotexs.ntu.edu.sg/files/ftp_upload/63509/63509fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table><tbody><tr><td>&lt;strong&gt;Equipment and supplies:&lt;/strong&gt;</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&#039;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 &amp;amp; 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 &amp;deg;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>&lt;strong&gt;Microscopes:&lt;/strong&gt;</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.&lt;br/&gt; Microscope control and image capture software (NIS-Elements by Nikon) is used here but any confocal microscope will work.</td></tr><tr><td>&lt;strong&gt;Reagents for surgeries and dissections:&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Calcium chloride dihydrate</td><td>Sigma</td><td>C7902</td><td>For Marc&#039;s Modified Ringers (MMR) and embryo medium (E3).</td></tr><tr><td>HEPES</td><td>Sigma</td><td>H7006</td><td>For Marc&#039;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&#039;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&#039;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&#039;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&#039;s Modified Ringers (MMR) and embryo medium (E3).</td></tr><tr><td>Sodium chloride</td><td>Sigma</td><td>S9888</td><td>For Marc&#039;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>&lt;strong&gt;Reagents for immunohistochemistry:&lt;/strong&gt;</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 &amp;deg;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 &amp;micro;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>&lt;strong&gt;Software for data analysis&lt;/strong&gt;</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>&lt;strong&gt;Zebrafish strains.&lt;/strong&gt; This study used the AB, TU, &lt;i&gt;Tg[atoh7:RFP]&lt;/i&gt; strains.</td><td></td><td></td><td>Available from the&amp;nbsp; 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 axon tracing with lipophilic DiI or DiO dyes25,26 or multispectral labeling with brainbow27, would also be possible.
As shown in Figure 2, progressive degeneration of retinal axons in the optic tectum neuropil was observed following eye removal. By two days post surgery, RFP-labeled axons exhibited hallmarks of rapid Wallerian degeneration28 such as blebbing and fragmentation (Figure 2A,B). Recent work has shown that microglia engulf more myelin material when neurons are silenced29. Consistent with microglia engulfing bits of the degenerating RGC axons and migrating away from the source of degeneration, bright RFP-labeled puncta within and outside the tectal neuropil were noted (Figure 1B, arrows). By four days post surgery, fragmented axons and RFP-labeled axonal debris are considerably reduced in either tectal lobe, indicating relatively rapid clearance of the dying and degenerating axons (Figure 2C,D).
To perform wholemount immunohistochemistry and visualize the optic tectum of larval zebrafish, the brain was exposed by dissecting off the eye(s), jaw, ears, and skull cap of the skin and connective tissues. Ideally, the brain remains intact during this procedure (e.g., Figure 2C,D). However, parts of the forebrain, particularly the olfactory bulb, are likely to be damaged or completely removed when the skull cap of skin or jaw is removed (Figure 2A, arrowhead). Moreover, sometimes the lateral edge of the tectum can be sliced when piercing or pinching the skin to pull it off the brain (e.g., Figure 2A; tear on the right tectal lobe).
Immunohistochemistry was also used to assess the number of mitotic cells in innervated and non-innervated lobes of the optic tectum by staining whole brains with a PH3 antibody. These data show significantly fewer mitotic cells on the denervated side of one-eyed larval fish (p = 0.00033, Welch&#39;s t-test; Figure 3).
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63509/63509fig02.jpg" /> 
Figure 2: Eye removal at 5 dpf triggers degeneration of optic nerve axons. (A-D)&#160;Representative maximum-intensity projections showing dorsal views of wild-type brains from intact larvae fixed at 7 dpf (A) or 9 dpf (C), or one-eyed larvae from eye extirpation surgeries at 5 dpf, fixed 2 dps (B) or 4 dps (D). All nuclei are stained with DAPI (green), and optic nerve axons terminating in the neuropil of the optic tectum are labeled by the atoh7:RFP transgene (magenta). Asterisk indicates innervated tectal neuropil in larvae with only the right eye intact. Arrows indicate examples of fragments of degenerated RGC axons emanating from the denervated optic tectum. Arrowhead indicates missing parts of the forebrain. Scale bar = 50 &#181;m. Abbreviations: dpf = days post fertilization; dps = days post surgery; RGC = retinal ganglion cells; DAPI = 4&#39;,6-diamidino-2-phenylindole. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63509/63509fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63509/63509fig03.jpg" /> 
Figure 3: Number of mitotic cells in the optic tectum increases with innervation. (A-B) Representative maximum-intensity projections showing dorsal views of wild-type brains from intact (A) or one-eyed (B) 9 dpf larvae immunostained with antibodies for the mitotic marker PH3 (magenta) and nuclei counterstained with DAPI (green). Asterisk indicates innervated tectal neuropil from the intact right eye. (C) Box plot with all individual data points overlaid showing quantitation of the ratio of PH3+ cells in innervated (left) versus denervated (right) tectal lobe as a difference ratio (left-right/total) for one-eyed (n = 12) and two-eyed (n = 8) 9 dpf larvae. Means of the two conditions compared with Welch&#39;s t-test (***, p &#60; 0.0005). Scale bar = 50 &#181;m. Abbreviations: dpf = days post fertilization; dps = days post surgery;&#160;PH3 = phosphorylated histone H3; DAPI = 4&#39;,6-diamidino-2-phenylindole. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63509/63509fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental file: Outline and instructions for plot shown in Figure 3C. This file contains all data and code so that anyone can reproduce the box plot with all data points overlaid, as shown in Figure 3C.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63509/JoVE-PH3-representative-plot_final_Rknit_revision.zip" target="_blank">Please click here to download this File.</a>

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

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

3.7K Views.  Reed College. Dus mijn lab is geïnteresseerd in het begrijpen hoe het oog en de hersenen samen groeien. Deze techniek stelt ons in staat om te bestuderen hoe retinale input de groei en ontwikkeling van de hersenen beïnvloedt. Chirurgische verwijdering van één oog van de levende larvale zebravis, gevolgd door observatie van het optische tectum stelt ons in staat om geïnnerveerde en gedenerveerde tectale kwabben binnen hetzelfde dier en tussen dieren te vergelijken.Het combineren van deze technie...

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

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 ...

VisioTracker, an Innovative Automated Approach to Oculomotor Analysis

Investigations into the visual system development and function necessitate quantifiable behavioral models of visual performance that are easy to elicit, robust, and simple to manipulate. A suitable model has been found in the optokinetic response (OKR), a reflexive behavior present in all vertebrates due to its high selection value. The OKR involves slow stimulus-following movements of eyes alternated with rapid resetting saccades. The measurement of this behavior is easily carried out in zebrafish larvae, due to its early and stable onset (fully developed after 96 hours post fertilization (hpf)), and benefitting from the thorough knowledge about zebrafish genetics, for decades one of the favored model organisms in this field. Meanwhile the analysis of similar mechanisms in adult fish has gained importance, particularly for pharmacological and toxicological applications.
Here we describe VisioTracker, a fully automated, high-throughput system for quantitative analysis of visual performance. The system is based on research carried out in the group of Prof. Stephan Neuhauss and was re-designed by TSE Systems. It consists of an immobilizing device for small fish monitored by a high-quality video camera equipped with a high-resolution zoom lens. The fish container is surrounded by a drum screen, upon which computer-generated stimulus patterns can be projected. Eye movements are recorded and automatically analyzed by the VisioTracker software package in real time.
Data analysis enables immediate recognition of parameters such as slow and fast phase duration, movement cycle frequency, slow-phase gain, visual acuity, and contrast sensitivity.
Typical results allow for example the rapid identification of visual system mutants that show no apparent alteration in wild type morphology, or the determination of quantitative effects of pharmacological or toxic and mutagenic agents on visual system performance.

The VisioTracker is an automated system for the quantitative analysis of visual performance of larval and small adult fish based on the recording of eye movements. It features full control over visual stimulus properties and real-time analysis, enabling high-throughput research in fields such as visual system development and function, pharmacology, neural circuit studies and sensorimotor integration.

Investigations into the visual system development and function necessitate quantifiable behavioral models of visual performance that are easy to elicit, ...

Neuroscience

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

Lens Transplantation in Zebrafish and its Application in the Analysis of Eye Mutants

The lens plays an important role in the development of the optic cup[1,2]. Using the zebrafish as a model organism, questions regarding lens development can be addressed. The zebrafish is useful for genetic studies due to several advantageous characteristics, including small size, high fecundity, short lifecycle, and ease of care. Lens development occurs rapidly in zebrafish. By 72 hpf, the zebrafish lens is functionally mature [3]. Abundant genetic and molecular resources are available to support research in zebrafish. In addition, the similarity of the zebrafish eye to those of other vertebrates provides basis for its use as an excellent animal model of human defects[4-7]. Several zebrafish mutants exhibit lens abnormalities, including high levels of cell death, which in some cases leads to a complete degeneration of lens tissues [8]. To determine whether lens abnormalities are due to intrinsic causes or to defective interactions with the surrounding tissues, transplantation of a mutant lens into a wild-type eye is performed. Using fire-polished metal needles, mutant or wild-type lenses are carefully dissected from the donor animal, and transferred into the host. To distinguish wild-type and mutant tissues, a transgenic line is used as the donor. This line expresses membrane-bound GFP in all tissues, including the lens. This transplantation technique is an essential tool in the studies of zebrafish lens mutants.

Lens development involves interactions with other tissues. Several zebrafish eye mutants are characterized by an abnormally small lens size. Here we demonstrate a lens transplantation experiment to determine whether this phenotype is due to intrinsic causes or defective interactions with tissues that surround the lens.

The lens plays an important role in the development of the optic cup[1,2]. Using the zebrafish as a model organism, questions regarding lens development ...

Visualization of the Superior Ocular Sulcus during Danio rerio Embryogenesis

Congenital ocular coloboma is a genetic disorder that is typically observed as a cleft in the inferior aspect of the eye resulting from incomplete choroid fissure closure. Recently, the identification of individuals with coloboma in the superior aspect of the iris, retina, and lens led to the discovery of a novel structure, referred to as the superior fissure or superior ocular sulcus (SOS), that is transiently present on the dorsal aspect of the optic cup during vertebrate eye development. Although this structure is conserved across mice, chick, fish, and newt, our current understanding of the SOS is limited. In order to elucidate factors that contribute to its formation and closure, it is imperative to be able to observe it and identify abnormalities, such as delay in the closure of the SOS. Here, we set out to create a standardized series of protocols that can be used to efficiently visualize the SOS by combining widely available microscopy techniques with common molecular biology techniques such as immunofluorescent staining and mRNA overexpression. While this set of protocols focuses on the ability to observe SOS closure delay, it is adaptable to the experimenter&#39;s needs and can be easily modified. Overall, we hope to create an approachable method through which our understanding of the SOS can be advanced to expand the current knowledge of vertebrate eye development.

Visualization of the Superior Ocular Sulcus during Danio rerio Embryogenesis

Here, we present a standardized series of protocols to observe the superior ocular sulcus, a recently-identified, evolutionarily-conserved structure in the vertebrate eye. Using zebrafish larvae, we demonstrate techniques necessary to identify factors that contribute to the formation and closure of the superior ocular sulcus.

Congenital ocular coloboma is a genetic disorder that is typically observed as a cleft in the inferior aspect of the eye resulting from incomplete choroid ...

Electroretinogram Recording in Larval Zebrafish using A Novel Cone-Shaped Sponge-tip Electrode

The zebrafish (Danio rerio) is commonly used as a vertebrate model in developmental studies and is particularly suitable for visual neuroscience. For functional measurements of visual performance, electroretinography (ERG) is an ideal non-invasive method, which has been well established in higher vertebrate species. This approach is increasingly being used for examining the visual function in zebrafish, including during the early developmental larval stages. However, the most commonly used recording electrode for larval zebrafish ERG to date is the glass micropipette electrode, which requires specialized equipment for its manufacture, presenting a challenge for laboratories with limited resources. Here, we present a larval zebrafish ERG protocol using a cone-shaped sponge-tip electrode. The novel electrode is easier to manufacture and handle, more economical, and less likely to damage the larval eye than the glass micropipette. Like previously published ERG methods, the current protocol can assess outer retinal function through photoreceptor and bipolar cell responses, the a- and b-wave, respectively. The protocol can clearly illustrate the refinement of visual function throughout the early development of zebrafish larvae, supporting the utility, sensitivity, and reliability of the novel electrode. The simplified electrode is particularly useful when establishing a new ERG system or modifying existing small-animal ERG apparatus for zebrafish measurement, aiding researchers in the visual neurosciences to use the zebrafish model organism.

Here, we present a protocol that simplifies the measurement of light evoked electroretinogram responses from larval zebrafish. A novel cone-shaped sponge-tip electrode can help to make the study of visual development in larval zebrafish using the electroretinogram ERG easier to achieve with reliable outcomes and lower cost.

The zebrafish (Danio rerio) is commonly used as a vertebrate model in developmental studies and is particularly suitable for visual neuroscience. For ...

Visualizing Ocular Morphogenesis by Lightsheet Microscopy Using rx3:GFP Transgenic Zebrafish

Vertebrate eye development is a complex process that begins near the end of embryo gastrulation and requires the precise coordination of cell migration, proliferation, and differentiation. Time-lapse imagining offers unique insight to the behavior of cells during eye development because it allows us to visualize oculogenesis in vivo. Zebrafish are an excellent model to visualize this process due to their highly conserved vertebrate eye and their ability to develop rapidly and externally while remaining optically transparent. Time-lapse imaging studies of zebrafish eye development are greatly facilitated by use of the transgenic zebrafish line Tg(rx3:GFP). In the developing forebrain, rx3:GFP expression marks the cells of the single eye field, and GFP continues to be expressed as the eye field evaginates to form an optic vesicle, which then invaginates to form an optic cup. High resolution time lapse imaging of rx3:GFP expression, therefore, allows us to track the eye primordium through time as it develops into the retina. Lightsheet microscopy is an ideal method to image ocular morphogenesis over time due to its ability to penetrate thicker samples for fluorescent imaging, minimize photobleaching and phototoxicity, and image at a high speed. Here, a&#160;protocol is&#160;provided&#160;for time-lapse imaging of ocular morphogenesis using a commercially available lightsheet microscope and an image processing workstation to analyze the resulting data. This protocol details the procedures for embryo anesthesia, embedding in low melting temperature agarose, suspension in the imaging chamber, setting up the imaging parameters, and finally analyzing the imaging data using image analysis software. The resulting dataset can provide valuable insights into the process of ocular morphogenesis, as well as perturbations to this process as a result of genetic mutation, exposure to pharmacological agents, or other experimental manipulations.

Here, a&#160;protocol is&#160;provided&#160;for time-lapse imaging of ocular morphogenesis using a commercially available lightsheet microscope and an image processing workstation to analyze the resulting data. This protocol details the procedures for embryo anesthesia, embedding in low melting temperature agarose, suspension in the imaging chamber, setting up the imaging parameters, and finally analyzing the imaging data using image analysis software.&#160;

Vertebrate eye development is a complex process that begins near the end of embryo gastrulation and requires the precise coordination of cell migration, ...

Electroretinogram Analysis of the Visual Response in Zebrafish Larvae

The electroretinogram (ERG) is a noninvasive electrophysiological method for determining retinal function. Through the placement of an electrode on the surface of the cornea, electrical activity generated in response to light can be measured and used to assess the activity of retinal cells in vivo. This manuscript describes the use of the ERG to measure visual function in zebrafish. Zebrafish have long been utilized as a model for vertebrate development due to the ease of gene suppression by morpholino oligonucleotides and pharmacological manipulation. At 5-10 dpf, only cones are functional in the larval retina. Therefore, the zebrafish, unlike other animals, is a powerful model system for the study of cone visual function in vivo. This protocol uses standard anesthesia, micromanipulation and stereomicroscopy protocols that are common in laboratories that perform zebrafish research. The outlined methods make use of standard electrophysiology equipment and a low light camera to guide the placement of the recording microelectrode onto the larval cornea. Finally, we demonstrate how a commercially available ERG stimulator/recorder originally designed for use with mice can easily be adapted for use with zebrafish. ERG of larval zebrafish provides an excellent method of assaying cone visual function in animals that have been modified by morpholino oligonucleotide injection as well as newer genome engineering techniques such as Zinc Finger Nucleases (ZFNs), Transcription Activator-Like Effector Nucleases (TALENs), and Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9, all of which have greatly increased the efficiency and efficacy of gene targeting in zebrafish. In addition, we take advantage of the ability of pharmacological agents to penetrate zebrafish larvae to evaluate the molecular components that contribute to the photoresponse. This protocol outlines a setup that can be modified and used by researchers with various experimental goals.

We present a method for the electroretinographic (ERG) analysis of zebrafish larvae utilizing micromanipulation and electroretinography techniques. This is a simple and straightforward method for assaying visual function of zebrafish larvae in vivo.

The electroretinogram (ERG) is a noninvasive electrophysiological method for determining retinal function. Through the placement of an electrode on the ...

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 ...

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

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