We thank Ralph Nelson and Daniel Possin for thoughtful guidance while developing this protocol, and Eva Ma, Ashley George and Gail Stanton for generation of stable transgenic zebrafish lines. The work was supported by NSF GRFP 2013158531 to M.G., NIH NEI 5T32EY007031 to W.C. and M.G., and EY026020 to J.H. and S.B.

All animal experiments were approved by the University of Washington Institutional Animal Care and Use Committee.
1. Preparing Animals and Equipment
NOTE: The retinal pigment epithelium (RPE) is a dark sheet of tissue surrounding the outside of the retina whose pigmentation can obscure retinal features and damage the tissue when confocal imaging ex vivo. In darkness, the RPE of zebrafish is retracted away from the retina; dark adapt fish to facilitate future removal of the RPE from the retina before slicing and imaging.
<ol>
	<li>Transfer fish to a spawning tank filled with fish water, and then wrap the spawning tank with dark fabric or place it in a dark cabinet.
	<ol>
		<li>Dark adapt zebrafish for at least 1 h prior to euthanasia to allow near complete separation of the RPE from the retina. 30 min dark adaptation is sufficient to remove most RPE, though pieces of it may remain intercalated between photoreceptors.</li>
	</ol>
	</li>
	<li>Melt ~ 15 mL of petroleum jelly in a 50-mL beaker on a hot plate and then draw 3 mL of the liquid into a 3-mL slip tip syringe. Invert syringe, place in a test tube rack, and allow petroleum jelly to cool.</li>
	<li>Make a reusable slicing chamber on a plain 7 cm X 2.5 cm microscope slide.
	<ol>
		<li>Using clear nail polish, paint narrow lines to create a 3 cm X 2.5 cm rectangle in the center of the slide, allow it to dry, and then add another layer of nail polish to the lines.</li>
	</ol>
	</li>
	<li>Prepare imaging ladders on cover slips to hold slices during imaging. The slices will form the &#34;rungs&#34; of the ladder.
	<ol>
		<li>For static imaging or injection experiments with minimal solution flow, make petroleum jelly ladders consisting of two flat wide parallel strips of petroleum jelly on 18 mm square glass cover slips. Use the syringe to apply two flattened ~ 1 cm long smears of cooled petroleum jelly 0.5 cm apart on the cover slip.</li>
	</ol>
	</li>
	<li>Ready the tissue slicer.
	<ol>
		<li>Clean a double edge razor blade with ethanol and allow it to air dry. Cut it into quarters with scissors, first lengthwise into halves then across each blade.</li>
		<li>Place the slicing chamber on the stage of the tissue slicer, center it horizontally on the stage and mark a long edge with permanent marker for alignment.</li>
		<li>Load a blade section onto the tissue slicer arm, ensure the blade lies flat and centered on the slide without touching the nail polish, then gently tighten the blade apparatus. Lower the blade arm by adjusting the knob &#188; turn, then place a scrap of filter paper in the center of the imaging chamber and test cut it. If the paper is not cut fully through, remount the blade.</li>
	</ol>
	</li>
	<li>Using the syringe, place a single small dot of cooled petroleum jelly in a 10-cm Petri dish ~ 1.5 cm to the right of the center. Press an imaging ladder into it using forceps with the petroleum jelly facing up. Make another small petroleum jelly dot ~ 1 cm from the inlet edge of the imaging chamber.</li>
	<li>Fit the petroleum jelly syringe with a 20g needle and uncap it. Hold the needle onto the syringe and use it to make two 1 cm long thin parallel strips of petroleum jelly lengthwise in the center of the slicing chamber. Space the strips ~ 1 cm apart.</li>
	<li>Make a reusable wire eye loop tool by wrapping the center of a ~ 4 cm segment of 30 g tungsten wire around a pair of closed forceps once tightly. Adjust the diameter of the loop by sliding the wire up or down the forceps until it is slightly larger than a zebrafish eye, typically ~ 2-3 mm. Twist the wire ends and secure them to the end of a 6-cm wooden stick using laboratory tape.</li>
	<li>Prepare Ringer&#39;s solution.
	<ol>
		<li>Thaw 50X supplement stock solution (Table 1) and add it fresh to HEPES-buffered, non-bicarbonate Ringer&#39;s solution (Table 2) the morning of the experiment; dilute 200 &#181;L of supplement stock solution in every 10 mL of Ringer&#39;s solution in a conical centrifuge tube or sterile glass bottle. For static imaging experiments, prepare at least 30 mL of Ringer&#39;s solution per experiment. The volume of Ringer&#39;s solution needed for perfusion experiments will depend on the flow rate and total experiment time.</li>
		<li>Check that the pH of the supplemented solution is 7.4 using a digital pH probe or pH paper, and adjust accordingly with dilute NaOH or HCl.</li>
		<li>Oxygenate supplemented Ringer&#39;s solution on ice by bubbling with 100% oxygen gas for at least 5 min using a standard medical oxygen tank and regulator fitted with a hose, or the optional gas bubbler manifold used for perfusion. Store oxygenated Ringer&#39;s solution on ice in a sealed conical centrifuge tube or sterile glass bottle near the dissection microscope; use this solution for dissection, imaging, and to dilute dyes or pharmacological agents.</li>
		<li>If other solutions are being used in the experiment, such as Na+-free Ringer&#39;s solution (Table 3), repeat steps 1.9.1.-1.9.3.</li>
	</ol>
	</li>
	<li>Gather Petri dishes, forceps, micro-scissors and other tools near the dissection microscope, and prepare a fish water ice bath for zebrafish euthanasia.</li>
</ol>
2. Preparing retinal slices (see Figure 1)
<ol>
	<li>Working under red ambient light to minimize light adaptation (which can make the RPE stick more tightly to the retina), euthanize zebrafish by immersion in the ice bath until touch response is lost, typically 1-2 min. Transfer the fish to a Petri dish and use a scalpel to cervically dislocate but not decapitate the fish.</li>
	<li>Use the wire loop to loosen connective tissue around one eye, and then pull the eye forward gently with the loop in one hand. Using micro-scissors in the other hand, cut the white optic nerve under the eye, taking care not to cut the back of the eye.</li>
	<li>Transfer the eye using forceps to a Petri dish of cold Ringer&#39;s solution on ice, and repeat for the second eye. Keep the eyes in darkness or under red light until the RPE is removed from the retina.</li>
	<li>Dissect the eyecups under a low power dissection microscope in a drop of cold Ringer&#39;s solution on a plain glass slide in a Petri dish.
	<ol>
		<li>Pierce the cornea with fine forceps, then gently remove pieces of clear, brittle cornea and silvery sclera with forceps or scissors. Remove and discard the lens and most of the sclera (see Figure 1A), and handle the isolated eyecup minimally.</li>
		<li>Pieces of fat, small bits of sclera, and black RPE can remain attached to the eyecup and removed from the retina later. If the retina separates from the RPE in step 2.4.1, proceed with the same steps using extra caution not to damage the delicate isolated retina.</li>
		<li>Position eyecup open side down (RPE up) on the slide, and cut into thirds or quarters with a fresh single edge razor blade in one motion (see Figure 1B). Discard pieces of tissue that are highly curved.</li>
	</ol>
	</li>
	<li>Flat mount retina on filter paper.
	<ol>
		<li>Wet a piece of filter paper with Ringer&#39;s solution and place it on the slide next to the eyecup pieces. Use flat forceps when handling the intact wet filter paper to avoid puncturing it. Add cold Ringer&#39;s solution to cover both the filter paper and tissue.</li>
		<li>Using forceps in each hand, carefully drag the filter paper underneath each eyecup piece with the RPE and photoreceptors facing up, i.e. with the back of the eye facing up. Position the eyecup pieces in a single line along the center of the filter paper (see Figure 1C). Handle the eyecup pieces gently with fine forceps only near an edge or corner.</li>
		<li>To help retinas adhere to the filter paper, place the wet filter paper on a dry paper towel for 3 s to wick moisture downward, but don&#39;t let the tissue become dry. Repeat until the eyecup pieces lie flat on the filter paper. Applying gentle suction to the underside of the filter paper helps flatten the retina, but this step is not essential.</li>
	</ol>
	</li>
	<li>If black sheets of RPE remain on the eyecup pieces, use fine forceps to gently peel it away starting from one corner (see Figure 1D) while the tissue is sitting in a drop of Ringer&#39;s solution. Should the retina lift off the filter paper, repeat the wicking step in 2.5.3. The retina may appear pink due to unbleached visual pigments.</li>
	<li>Repeat retina dissection and flat mounting steps in 2.4-2.6 for the second eye, if desired, keeping the first flat-mounted retina immersed in cold Ringer&#39;s solution. To streamline the slicing procedure, pieces of both retinas may be placed on one filter paper. Once the RPE has been removed, the protocol can be carried out under normal room light unless experiments necessitate darkness.</li>
	<li>Place the filter paper on a slide and trim it into a rectangle with a single edge razor blade, leaving ~ 0.5 cm of filter paper on either side of the line of retinas. Move the filter paper to the prepared slicing chamber, push the long filter paper edges into the thin petroleum jelly lines using forceps, and immerse the retinas in 3-4 drops of cold Ringer&#39;s solution. 
	NOTE: Some dyes, such as lipophilic dyes, are best loaded into flat mounts at this stage prior to slicing. These can be loaded, wicked away with a tissue, and washed in the slicing chamber. For instance, C12 558/568 BODIPY intensely stains retinal cell membranes and photoreceptor outer segments (see Figure 4A) when loaded at ~ 5 &#181;g/mL for 15 min at room temperature (typically 23-27 &#176;C), followed by a wash in excess Ringer&#39;s solution.</li>
	<li>Transfer the slicing chamber to the tissue slicer stage, position the long edge along the marked line, and secure the chamber ends to the stage with laboratory tape. Starting at one end, cut the retina and filter paper using firm, gentle pressure on the slicing arm. Check that the first slice was cut fully, then use the micrometer to cut ~ 400 &#181;m slices.</li>
	<li>Assemble the imaging ladder.
	<ol>
		<li>Place the slicing chamber with sliced retinal sections in the Petri dish from step 1.7 adjacent to the imaging ladder (see Figure 1E). Fill the dish with cold Ringer&#39;s solution to submerge its contents.</li>
		<li>Using forceps and keeping slices submerged, gently transfer strips of filter paper and retina to the ladder by sliding the Petri dish from left to right. Take care not to touch retinas directly. Rotate the slices 90&#176; and bury the filter paper edges in petroleum jelly.</li>
		<li>Finely position retinal slices in the ladder with forceps so that retinal layers are clearly visible under the dissection microscope (see Figure 1G). For slices on each end of the ladder, ensure that the tissue is facing inward toward the other slices to minimize motion of the tissue during injection or flow. Discard any retinal slices that are not well adhered to the filter paper (see Figure 2A).</li>
	</ol>
	</li>
	<li>If desired, load dyes into retinal slices at this stage and wash in excess Ringer&#39;s solution prior to imaging. 
	NOTE: Propidium iodide (PI) and Hoechst 33342 robustly stain nuclei of dead and all cells, respectively (see Figure 2B), when incubated with retinal slices at 5 &#181;g/mL for 20 min at room temperature. Tetramethylrhodamine (TMRM) accumulates in actively respiring mitochondria throughout the retina when incubated at 1 nM for 30 min at room temperature.</li>
	<li>While the slices are staining, prepare the imaging chamber and injection apparatus. Use the syringe to flush the tubing with Ringer&#39;s solution and purge bubbles, attach the open end of the tubing to the imaging chamber inlet, and close the stopcock. Remove the syringe and fill it with reagent(s) for injection, then reattach it to the tubing.</li>
	<li>Use forceps to transfer the coverslip with retinal slices to the imaging chamber, pressing the coverslip into the dot of petroleum jelly near the inlet edge of the imaging chamber (see Figure 1H). Fill the imaging chamber with Ringer&#39;s solution to cover slices.</li>
</ol>
3. Imaging retinal slices
<ol>
	<li>Place the filled imaging chamber with the connected injection apparatus on the stage of an upright confocal microscope equipped with a 20 or 40X water dipping lens. Secure the imaging chamber with stage clips.</li>
	<li>Lower the dipping lens over the ladder, and focus on a slice at one end of the ladder under dim trans-illuminated light. Examine each slice for transverse orientation, presence of photoreceptor outer segments, and secure adhesion to the filter paper.</li>
	<li>Select the best slice for time lapse imaging and configure the microscope software for time lapse image acquisition. Table 4 outlines typical imaging settings for various fluorescent markers and dyes. 
	NOTE: The rate of image acquisition will vary depending on the microscope, fluorescent marker(s), and biological process being studied. For instance, use 800x800 pixel resolution, 2 &#181;s/pixel scan speed, and a 10-s frame rate for imaging calcium dynamics in photoreceptors with GCaMP3 and a red dye.
	<ol>
		<li>If available, use the software to trace physical landmarks in the slice (photoreceptor outer segments, cell bodies, nuclei) to aid potential reorientation during the time lapse. Also set up the software to monitor real-time fluorescence across the slice.</li>
	</ol>
	</li>
	<li>Begin imaging and monitor baseline fluorescence. When fluorescence across the slice stabilizes (typically within 2-5 min), proceed with the experiment. For injection, open the syringe stopcock, then slowly depress and draw back on the plunger twice to aid mixing. 
	NOTE: For instance, abolish mitochondrial function by injecting a concentrated solution of the protonophore CCCP to reach a final concentration of 1 &#181;M in the imaging chamber. This induces a robust Ca2+ burst in photoreceptor cytosol reported by GCaMP, then a steady decrease in mitochondrial membrane potential reported by TMRM.</li>
	<li>Closely monitor slices for drift during the experiment, and use the software to make micro-adjustments according to physical landmarks. Typically drift in the Z-direction during injection or perfusion is &#60; 5 &#181;m.</li>
</ol>
4. Imaging retinal slices during perfusion experiments where solutions are changed or flowed continuously
NOTE: Imaging retinal slices during perfusion experiments where solutions are changed or flowed continuously is similar to setup for static imaging or injection experiments, with the following modifications.
<ol>
	<li>Instead of petroleum jelly ladders described in step 1.4, use sturdier wax ladders to hold retinal slices steady during solution flow.
	<ol>
		<li>Place two small parallel cylinders of unflavored dental wax ~ 0.5 cm apart on a coverslip. On a flat surface, use a thumb to press each cylinder down and out toward the parallel edge of the coverslip. Score both flattened cylinders horizontally with a #1 coverslip (see Figure 1E, right side) then smear a thin layer of petroleum jelly between the wax strips with a spatula.</li>
		<li>When assembling the imaging ladder in step 2.10.2, press the sliced filter paper edges into the wax scores using fine forceps (Figure 1G).</li>
	</ol>
	</li>
	<li>To set up for perfusion in step 2.12, fill syringe reservoirs with preoxygenated solutions, or use the optional gas manifold to oxygenate solutions in each reservoir. Flush all tubing with Ringer&#39;s solution, ensure all lines flow when opened and purge large bubbles.</li>
	<li>Before filling the imaging chamber in step 2.13, use the syringe to place another dot of petroleum jelly over each exposed corner of the coverslip to prevent it from sliding laterally during flow.</li>
	<li>When the imaging chamber is filled and mounted on the microscope stage, turn on the aspirator and connect the aspirator tubing. Test the flow of the Ringer&#39;s solution and use the micropositioner to situate the aspirator tube over the outflow chamber so that small amounts of liquid are drawn off before the chamber overflows (typically ~ 1 mm above the solution surface for a 2 mL/min flow rate). Keep Ringer&#39;s solution flowing while selecting slices in step 3.4.</li>
	<li>To conduct the experiment in step 3.4 for perfusion, switch flow of solutions by closing the stopcock of the first solution while opening that of the second solution.
	<ol>
		<li>For instance, to deplete extracellular Na+ from the imaging chamber, use two syringe reservoirs filled with Ringer&#39;s solution or Na+-free solution (Table 3). Flow Ringer&#39;s solution to establish baseline, then switch to Na+-free solution. This results in large cytosolic Ca2+ increases in photoreceptor outer segments and cell bodies (see Figure 4).</li>
	</ol>
	</li>
	<li>For gravity-fed perfusion systems, monitor the level of solution in each reservoir so the flow rate is constant during imaging. Top off reservoirs as needed with oxygenated solutions, or maintain continuous oxygen bubbling with the gas manifold.</li>
</ol>

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The authors declare that they have no competing financial interests.

Ex vivo imaging of fresh zebrafish retinal slices has proven to be a versatile tool for studying photoreceptor biology20,21,22, and is unique in that it enables analysis of single cells in a mature, fully differentiated retina. With practice, it is possible to conduct multiple experiments with tissue from a single fish, even using serial slices from the same part of the retina. In addition to the challenges and suggesti...

The zebrafish (Danio rerio) has become widely used in medical and basic scientific research1, owing to its small size, rapid development and vertebrate organ systems. The natural transparency of zebrafish larvae combined with established methods for transgenesis have enabled detailed visualization of cellular processes in a living animal. A number of genetically encoded fluorescent biosensors have been targeted to specific zebrafish cells to detect Ca2+ 2, hydrogen peroxide3, apoptotic activation4 and ATP5.
In vivo imaging of zebrafish larvae has led to breakthroughs in the field of neuroscience, including mapping of brain circuitry6 and drug development for central nervous system disorders7. Zebrafish are well suited for vision research because their retinas feature the laminar structure and neuron types of higher vertebrates, and they display robust visual behaviors8,9. Several types of retinal degenerations analogous to human disease have been modeled successfully and studied in zebrafish10,11, including live imaging of individual photoreceptors degenerating within a retina2,12.
While in vivo larval zebrafish imaging is a valuable tool, it becomes more challenging as fish grow and develop pigmentation, and some pharmacological treatments cannot permeate an entire animal. Further, certain cellular processes change with development and age, making later time points critical for understanding function and the progression of disease in adult animals. Biochemical methods such as immunoblot, quantitiative PCR, O2 consumption, and metabolomic analyses can provide important clues about biology of the retina as a whole, but it is difficult to discern contributions of individual cell types affected by disease. Imaging isolated retinal tissue ex vivo bypasses these issues, and while imaging flat mounted retinas affords a view of the outer retina13, deeper inner retinal features are obscured. Transverse retinal slices, such as those presented in fixed immunohistochemical analyses, enable a clear view of all layers and cell types but only offer a single snapshot of the dynamic processes involved in normal function and disease.
Here, we present a method for generating ex vivo transverse retinal slices from adult zebrafish for imaging. It is similar to methods for preparing amphibian and zebrafish retinal slices for electrophysiological and morphological studies14,15, with important modifications for time lapse imaging ex vivo using confocal microscopy. Fluorescence responses of biosensors or dyes in slices are monitored in real time with a confocal microscope while delivering pharmacological agents using perfusion. While the method was developed for imaging photoreceptors, it may be feasible to use it for visualizing M&#252;ller cells, bipolar cells, horizontal cells, amacrine cells, or retinal ganglion cells with appropriate fluorescent markers. Additionally, slices can be loaded with fluorescent cell-permeable dyes to report cell viability, vesicular transport, mitochondrial function, or redox state. This versatile preparation allows visualization of a wide range of subcellular processes throughout the retina, including Ca2+ dynamics, signal transduction and metabolic state.

<table><tbody><tr><td>zebrafish</td><td>Univeristy of Washington South Lake Union Aquatics Facility</td><td></td><td>stocks maintained in-house as stable transgenic lines</td></tr><tr><td>petroleum jelly</td><td>Fisher Scientific</td><td>19-090-843</td><td>for petroleum jelly syringe</td></tr><tr><td>3-mL slip tip syringe</td><td>Fisher Scientific</td><td>14-823-436</td><td>for petroleum jelly syringe</td></tr><tr><td>20g 3.8 cm slip tip needle</td><td>Fisher Scientific</td><td>14-826-5B</td><td>for petroleum jelly syringe</td></tr><tr><td>plain 7 cm X 2.5 cm microscope slide</td><td>Fisher Scientific</td><td>12-550-A3</td><td>for eyecup dissection, slicing chamber</td></tr><tr><td>Seche Vite clear nail polish</td><td>Amazon</td><td>B00150LT40</td><td>for slicing chamber</td></tr><tr><td>18 mm X 18 mm #1 glass coverslips</td><td>Fisher Scientific</td><td>12-542A</td><td>for imaging ladders</td></tr><tr><td>unflavored dental wax</td><td>Amazon</td><td>B01K8WNL5A</td><td>for imaging ladders</td></tr><tr><td>double edge razor blades</td><td>Stoelting</td><td>51427</td><td>for tissue slicing</td></tr><tr><td>tissue slicer with digital micrometer</td><td>Stoelting</td><td>51415</td><td>for tissue slicing</td></tr><tr><td>filter paper - white gridded mixed cellulose, 13 mm diameter, 0.45 &amp;micro;m pore size</td><td>EMD Millipore</td><td>HAWG01300</td><td>filter paper for mounting retinas</td></tr><tr><td>10 cm petri dish</td><td>Fisher Scientific</td><td>FB0875712</td><td>for fish euthanasia, dissection, imaging ladder assembly</td></tr><tr><td>15 cm plain-tipped wood applicator stick</td><td>Fisher Scientific</td><td>23-400-112</td><td>for wire eye loop tool</td></tr><tr><td>30g (0.25 mm diameter) tungsten wire</td><td>Fisher Scientific</td><td>AA10408G6</td><td>for wire eye loop tool</td></tr><tr><td>D-glucose</td><td>Sigma Aldrich</td><td>G8270</td><td>component of supplement stock solution</td></tr><tr><td>sodium L-lactate</td><td>Sigma Aldrich</td><td>L7022</td><td>component of supplement stock solution</td></tr><tr><td>sodium pyruvate</td><td>Sigma Aldrich</td><td>P2256</td><td>component of supplement stock solution</td></tr><tr><td>L-glutamine</td><td>Sigma Aldrich</td><td>G3126</td><td>component of supplement stock solution</td></tr><tr><td>&amp;nbsp;L-glutathione, reduced</td><td>Sigma Aldrich</td><td>G4251</td><td>component of supplement stock solution</td></tr><tr><td>L-ascorbic acid</td><td>Sigma Aldrich</td><td>A5960</td><td>component of supplement stock solution</td></tr><tr><td>NaCl</td><td>Sigma Aldrich</td><td>S7653</td><td>component of Ringer&#039;s solution</td></tr><tr><td>KCl</td><td>Sigma Aldrich</td><td>P9333</td><td>component of Ringer&#039;s solution</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt; &amp;middot; 2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma Aldrich</td><td>C3881</td><td>component of Ringer&#039;s solution</td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma Aldrich</td><td>S8282</td><td>component of Ringer&#039;s solution</td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt; &amp;middot; 6H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma Aldrich</td><td>M0250</td><td>component of Ringer&#039;s solution</td></tr><tr><td>HEPES</td><td>Sigma Aldrich</td><td>H3375</td><td>component of Ringer&#039;s solution</td></tr><tr><td>Tris base</td><td>Fisher Scientific</td><td>BP152</td><td>component of Na&lt;sup&gt;+&lt;/sup&gt;-free Ringer&#039;s solution</td></tr><tr><td>6 N HCl</td><td>Fisher Scientific</td><td>02-003-063</td><td>component of Na&lt;sup&gt;+&lt;/sup&gt;-free Ringer&#039;s solution</td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma Aldrich</td><td>P5655</td><td>component of Na&lt;sup&gt;+&lt;/sup&gt;-free Ringer&#039;s solution</td></tr><tr><td>50 mL conical centrifuge tube</td><td>Denville Scientific</td><td>C1062-P</td><td>container for Ringer&#039;s solution</td></tr><tr><td>Vannas scissors - 8 cm, angled 5 mm blades</td><td>World Precision Instruments</td><td>501790</td><td>micro-scissors for eyecup dissection</td></tr><tr><td>Swiss tweezers - #5, 11 cm, straight, 0.06 X 0.07 mm tips</td><td>World Precision Instruments</td><td>504510</td><td>fine forceps for eyecup dissection and slice manipulation</td></tr><tr><td>single edge razor blades</td><td>Fisher Scientific</td><td>12-640</td><td>for eyecup dissection and trimming filter paper</td></tr><tr><td>EMD Millipore filter forceps</td><td>Fisher Scientific</td><td>XX6200006P</td><td>flat forceps for handling wet filter paper</td></tr><tr><td>C&lt;sup&gt;12&lt;/sup&gt; 558/568 BODIPY</td><td>Fisher Scientific</td><td>D3835</td><td>stains live cell nuclei; incubate 5 &amp;micro;g/mL for 15 min at room temperature</td></tr><tr><td>propidium iodide (PI)</td><td>Fisher Scientific</td><td>P3566</td><td>stains dead cell nuclei; incubate 5 &amp;micro;g/mL for 20 min at room temperature</td></tr><tr><td>Hoechst 33342</td><td>Fisher Scientific</td><td>62249</td><td>stains live cell nuclei; incubate 5 &amp;micro;g/mL for 20 min at room temperature</td></tr><tr><td>Tetramethylrhodamine, methyl ester (TMRM)</td><td>Fisher Scientific</td><td>T668</td><td>stains functional, negatively-charged mitochondria; incubate 1 nM for 30 min at room temperature</td></tr><tr><td>tissue perfusion chamber</td><td>Cell MicroControls</td><td>BT-1-18/BT-1-18BV [-SY]</td><td>imaging chamber for injection or perfusion</td></tr><tr><td>2-(N-(7-Nitrobenz-2-oxa-1,3-diazol-4-yl)Amino)-2-Deoxyglucose (NBDG)</td><td>Fisher Scientific</td><td>N13195</td><td>fluorescent glucose analog adminitered orally to zebrafish 30 min prior to euthanasia</td></tr><tr><td>Olympus laser scanning confocal microscope</td><td>Olympus</td><td>FV1000</td><td>confocal microscope for visualizing fluorescence of slices at single-cell resolution</td></tr><tr><td>Carbonyl cyanide 3-chlorophenylhydrazone (CCCP)</td><td>Sigma Aldrich</td><td>C2759</td><td>experimental reagent which ablates mitochondrial respiration; treat slices to a final concentration of 1 &amp;micro;M</td></tr><tr><td>miniature aspirator positioner</td><td>Cell MicroControls</td><td>FL-1</td><td>for perfusion</td></tr><tr><td>perfusion manifold, gas bubbler manifold, flow valve, 60cc syringe holder</td><td>Warner Instruments</td><td>various</td><td>for perfusion</td></tr></tbody></table>

preparing fresh retinal slices from adult zebrafish for ex vivo imaging experiments

The retina is a complex tissue that initiates and integrates the first steps of vision. Dysfunction of retinal cells is a hallmark of many blinding diseases, and future therapies hinge on fundamental understandings about how different retinal cells function normally. Gaining such information with biochemical methods has proven difficult because contributions of particular cell types are diminished in the retinal cell milieu. Live retinal imaging can provide a view of numerous biological processes on a subcellular level, thanks to a growing number of genetically encoded fluorescent biosensors. However, this technique has thus far been limited to tadpoles and zebrafish larvae, the outermost retinal layers of isolated retinas, or lower resolution imaging of retinas in live animals. Here we present a method for generating live ex vivo retinal slices from adult zebrafish for live imaging via confocal microscopy. This preparation yields transverse slices with all retinal layers and most cell types visible for performing confocal imaging experiments using perfusion. Transgenic zebrafish expressing fluorescent proteins or biosensors in specific retinal cell types or organelles are used to extract single-cell information from an intact retina. Additionally, retinal slices can be loaded with fluorescent indicator dyes, adding to the method&#39;s versatility. This protocol was developed for imaging Ca2+ within zebrafish cone photoreceptors, but with proper markers it could be adapted to measure Ca2+ or metabolites in M&#252;ller cells, bipolar and horizontal cells, microglia, amacrine cells, or retinal ganglion cells. The retinal pigment epithelium is removed from slices so this method is not suitable for studying that cell type. With practice, it is possible to generate serial slices from one animal for multiple experiments. This adaptable technique provides a powerful tool for answering many questions about retinal cell biology, Ca2+, and energy homeostasis.

Stable positioning and transverse orientation of slices are key to successful imaging with injection or perfusion of pharmacological agents. Carefully examine and reposition slices prior to confocal imaging as needed to ensure all retinal layers are visible (Figure 2A, slice ii). If a slice is rotated slightly forward (Figure 2A, slice iii), bundles of outer segments will be visible and small adjustments can be made with forceps ...

Watch this Scientific Journal Video about Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments at JoVE.com

Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments

Imaging retinal tissue can provide single-cell information that cannot be gathered from traditional biochemical methods. This protocol describes preparation of retinal slices from zebrafish for confocal imaging. Fluorescent genetically encoded sensors or indicator dyes allow visualization of numerous biological processes in distinct retinal cell types.

Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments

The retina is a complex tissue that initiates and integrates the first steps of vision. Dysfunction of retinal cells is a hallmark of many blinding ...

preparing-fresh-retinal-slices-from-adult-zebrafish-for-ex-vivo

Research

JoVE Journal

Neuroscience

9.1K Views.  University of Washington. Imaging retinal tissue can provide single-cell information that cannot be gathered from traditional biochemical methods. This protocol describes preparation of retinal slices from zebrafish for confocal imaging. Fluorescent genetically encoded sensors or indicator dyes allow visualization of numerous biological processes in distinct retinal cell types.

Video: Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments

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

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

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

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

Developmental Biology

Correlative Super-resolution and Electron Microscopy to Resolve Protein Localization in Zebrafish Retina

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and scanning electron microscopy. The sub-diffraction limit resolution capabilities of super-resolution light microscopes allow improving the accuracy of the correlated data. Briefly, 110 nanometer thick cryo-sections are transferred to a silicon wafer and, after immunofluorescence staining, are imaged by super-resolution light microscopy. Subsequently, the sections are preserved in methylcellulose and platinum shadowed prior to imaging in a scanning electron microscope (SEM). The images from these two microscopy modalities are easily merged using tissue landmarks with open source software. Here we describe the adapted method for the larval zebrafish retina. However, this method is also applicable to other types of tissues and organisms. We demonstrate that the complementary information obtained by this correlation is able to resolve the expression of mitochondrial proteins in relation with the membranes and cristae of mitochondria as well as to other compartments of the cell.

This protocol describes the necessary steps to obtain subcellular protein localization results on zebrafish retina by correlating super-resolution light microscopy and scanning electron microscopy images.

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and ...

Retrograde Labeling of Retinal Ganglion Cells in Adult Zebrafish with Fluorescent Dyes

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs survival and regeneration experiments. Many studies have been performed in mammalian animals to research RGCs survival after optic nerve injury. However, retrograde labeling of RGCs in adult zebrafish has not yet been reported, though some alternative methods can count cell numbers in retinal ganglion cell layers (RGCL). Considering the small size of the adult zebrafish skull and the high risk of death after drilling on the skull, we open the skull with the help of acid-etching and seal the hole with a light curing bond, which could significantly improve the survival rate. After absorbing the dyes for 5 days, almost all the RGCs are labeled. As this method does not need to transect the optic nerve, it is irreplaceable in the research of RGCs survival after optic nerve crush in adult zebrafish. Here, we introduce this method step by step and provide representative results.

We introduce an efficient method to retrograde label retinal ganglion cells (RGCs) in adult zebrafish.

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs ...

A Novel Light Damage Paradigm for Use in Retinal Regeneration Studies in Adult Zebrafish

Light-induced retinal degeneration (LIRD) is commonly used in both rodents and zebrafish to damage rod and cone photoreceptors. In adult zebrafish, photoreceptor degeneration triggers M&#252;ller glial cells to re-enter the cell cycle and produce transient-amplifying progenitors. These progenitors continue to proliferate as they migrate to the damaged area, where they ultimately give rise to new photoreceptors. Currently, there are two widely-used LIRD paradigms, each of which results in varying degrees of photoreceptor loss and corresponding differences in the regeneration response. As more genetic and pharmacological tools are available to test the role of individual genes of interest during regeneration, there is a need to develop a robust LIRD paradigm. Here we describe a LIRD protocol that results in widespread and consistent loss of both rod and cone photoreceptors in which we have combined the use of two previously established LIRD techniques. Furthermore, this protocol can be extended for use in pigmented animals, which eliminates the need to maintain transgenic lines of interest on the albino background for LIRD studies.

Multiple light damage protocols have been described to damage photoreceptors and consequently induce a retinal regeneration response in adult zebrafish. This protocol describes an improved method that can be used in pigmented animals and that damages the vast majority of rod and cone photoreceptors across the entire retina.

Light-induced retinal degeneration (LIRD) is commonly used in both rodents and zebrafish to damage rod and cone photoreceptors. In adult zebrafish, ...

Microdissection of Zebrafish Embryonic Eye Tissues

Zebrafish is a popular animal model for research on eye development because of its rapid ex utero development and good fecundity. By 3 days post fertilization (dpf), the larvae will show the first visual response. Many genes have been identified to control a proper eye development, but we are far from a complete understanding of the underlying genetic architecture. Whole genome gene expression profiling is a useful tool to elucidate genetic regulatory network for eye development. However, the small size of the embryonic eye in zebrafish makes it challenging to obtain intact and pure eye tissues for expression analysis. For example, the anterior-posterior length of the eye between day 2 and 3 is only approximately 200-300 &mu;m, while the diameter of the lens is less 100 &mu;m. Also, the retinal pigment epithelium (RPE) underlying the retina is just a single-layer epithelium. While gene expression profiles can be obtained from the whole embryo, they do not accurately represent the expression of these tissues. Therefore pure tissue must be obtained for a successful gene expression profiling of eye development. To address this issue, we have developed an approach to microdissect intact retina and retina with RPE attached from 1-3 dpf, which cover major stages of eye morphogenesis. All procedures can be done with fine forceps and general laboratory supplies under standard stereomicroscopes. For retinal dissection, the single-layer RPE is removed and peeled off by brushing action and the preferential adherence of the RPE remnants to the surface of the culture plate for dissection. For RPE-attached retinal dissection, the adherence of RPE to the dissection plate is removed before the dissection so that the RPE can be completely preserved with the retina. A careful lifting action of this tissue can efficiently separate the presumptive choroid and sclera. The lens can be removed in both cases by a chemically etched tungsten needle. In short, our approach can obtain intact eye tissues and has been successfully utilized to study tissue-specific expression profiles of zebrafish retina1, 2 and retinal pigment epithelium3.

This article describes an approach to microdissect zebrafish retinas with and without retinal pigment epithelium attached, from one to three days postfertilization embryos.

Zebrafish is a popular animal model for research on eye development because of its rapid ex utero development and good fecundity. By 3 days post ...

Biology

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

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

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

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

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.

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 ...

Studying Diabetes Through the Eyes of a Fish: Microdissection, Visualization, and Analysis of the Adult tg(fli:EGFP) Zebrafish Retinal Vasculature

Diabetic retinopathy is the leading cause of blindness among middle-aged adults. The rising prevalence of diabetes worldwide will make the prevention of diabetic microvascular complications one of the key research fields of the next decades. Specialized, targeted therapy and novel therapeutic drugs are needed to manage the increasing number of patients at risk of vision-loss. The zebrafish is an established animal model for developmental research questions with increasing relevance for modeling metabolic multifactorial disease processes. The advantages of the species allow for optimal visualization and high throughput drug screening approaches, combined with the strong ability to knock out genes of interest. Here, we describe a protocol which will allow easy analysis of the adult tg(fli:EGFP) zebrafish retinal vasculature as a fast read-out in settings of long-term vascular pathologies linked to neoangiogenesis or vessel damage. This is achieved via dissection of the zebrafish retina and whole-mounting of the tissue. Visualization of the exposed vessels is then achieved via confocal microscopy of the green EGFP reporter expressed in the adult retinal vasculature. Correct handling of the tissue will lead to better outcomes and less internal vessel breakage to assure the visualization of the unaltered vascular structure. The method can be utilized in zebrafish models of retinal vasculopathy linked to changes in the vessel architecture as well as neoangiogenesis.

Studying Diabetes Through the Eyes of a Fish: Microdissection, Visualization, and Analysis of the Adult tgfli:EGFP Zebrafish Retinal Vasculature

Here, we discuss a method protocol which will allow an easy analysis of the adult tg(fli:EGFP) zebrafish retinal vasculature as a fast read-out in settings of long-term vascular pathologies linked to neoangiogenesis and structural changes.

Diabetic retinopathy is the leading cause of blindness among middle-aged adults. The rising prevalence of diabetes worldwide will make the prevention of ...

Medicine

ROS Live Cell Imaging During Neuronal Development

Reactive oxygen species (ROS) are well-established signaling molecules, which are important in normal development, homeostasis, and physiology. Among the different ROS, hydrogen peroxide (H2O2) is best characterized with respect to roles in cellular signaling. H2O2 has been implicated during the development in several species. For example, a transient increase in H2O2 has been detected in zebrafish embryos during the first days following fertilization. Furthermore, depleting an important cellular H2O2 source, NADPH oxidase (NOX), impairs nervous system development such as the differentiation, axonal growth, and guidance of retinal ganglion cells (RGCs) both in&#160;vivo and in vitro. Here, we describe a method for imaging intracellular H2O2 levels in cultured zebrafish neurons and whole larvae during development using the genetically encoded H2O2-specific biosensor, roGFP2-Orp1. This probe can be transiently or stably expressed in zebrafish larvae. Furthermore, the ratiometric readout diminishes the probability of detecting artifacts due to differential gene expression or volume effects. First, we demonstrate how to isolate and culture RGCs derived from zebrafish embryos that transiently express roGFP2-Orp1. Then, we use whole larvae to monitor H2O2 levels at the tissue level. The sensor has been validated by the addition of H2O2. Additionally, this methodology could be used to measure H2O2 levels in specific cell types and tissues by generating transgenic animals with tissue-specific biosensor expression. As zebrafish facilitate genetic and developmental manipulations, the approach demonstrated here could serve as a pipeline to test the role of H2O2 during neuronal and general embryonic development in vertebrates.

This protocol describes the use of a genetically encoded hydrogen peroxide (H2O2)-biosensor in cultured zebrafish neurons and larvae for assessing the physiological signaling roles of H2O2 during nervous system development. It can be applied to different cell types and modified with experimental treatments to study reactive oxygen species (ROS) in general development.

Reactive oxygen species (ROS) are well-established signaling molecules, which are important in normal development, homeostasis, and physiology. Among the ...

Nitroreductase/Metronidazole-Mediated Ablation and a MATLAB Platform (RpEGEN) for Studying Regeneration of the Zebrafish Retinal Pigment Epithelium

The retinal pigment epithelium (RPE) resides at the back of the eye and performs functions essential for maintaining the health and integrity of adjacent retinal and vascular tissues. At present, the limited reparative capacity of mammalian RPE, which is restricted to small injuries, has hindered progress to understanding in vivo RPE regenerative processes. Here, a detailed methodology is provided to facilitate the study of in vivo RPE repair utilizing the zebrafish, a&#160;vertebrate model capable of robust tissue regeneration. This protocol describes a transgenic nitroreductase/metronidazole (NTR/MTZ)-mediated injury paradigm (rpe65a:nfsB-eGFP), which results in ablation of the central two-thirds of the RPE after 24 h treatment with MTZ, with subsequent tissue recovery. Focus is placed on RPE ablations in larval zebrafish and methods for testing the effects of pharmacological compounds on RPE regeneration are also outlined. Generation and validation of RpEGEN, a MATLAB script created to automate quantification of RPE regeneration based on pigmentation, is also discussed. Beyond active RPE repair mechanisms, this protocol can be expanded to studies of RPE degeneration and injury responses as well as the effects of RPE damage on adjacent retinal and vascular tissues, among other cellular and molecular processes. This zebrafish system holds significant promise in identifying genes, networks, and processes that drive RPE regeneration and RPE disease-related mechanisms, with the long-term goal of applying this knowledge to mammalian systems and, ultimately, toward therapeutic development.

Nitroreductase/Metronidazole-Mediated Ablation and a MATLAB Platform RpEGEN for Studying Regeneration of the Zebrafish Retinal Pigment Epithelium

This protocol describes the methodology to genetically ablate the retinal pigment epithelium (RPE) using a transgenic zebrafish model. Adapting the protocol to incorporate signaling pathway modulation using pharmacological compounds is extensively detailed. A MATLAB platform for quantifying RPE regeneration based on pigmentation was developed and is presented and discussed.

The retinal pigment epithelium (RPE) resides at the back of the eye and performs functions essential for maintaining the health and integrity of adjacent ...

Preparing Fresh Retinal Slices from Adult Zebrafish for Ex Vivo Imaging Experiments

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Rozdziały w tym wideo

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

Correlative Super-resolution and Electron Microscopy to Resolve Protein Localization in Zebrafish Retina

Retrograde Labeling of Retinal Ganglion Cells in Adult Zebrafish with Fluorescent Dyes

A Novel Light Damage Paradigm for Use in Retinal Regeneration Studies in Adult Zebrafish

Microdissection of Zebrafish Embryonic Eye Tissues

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

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

Studying Diabetes Through the Eyes of a Fish: Microdissection, Visualization, and Analysis of the Adult tg(fli:EGFP) Zebrafish Retinal Vasculature

ROS Live Cell Imaging During Neuronal Development

Nitroreductase/Metronidazole-Mediated Ablation and a MATLAB Platform (RpEGEN) for Studying Regeneration of the Zebrafish Retinal Pigment Epithelium