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 border="0" cellpadding="0" cellspacing="0" width="1153">
	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:332px;">zebrafish</td>
			<td style="width:239px;">Univeristy of Washington South Lake Union Aquatics Facility</td>
			<td style="width:181px;"></td>
			<td style="width:401px;">stocks maintained in-house as stable transgenic lines</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">petroleum jelly</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">19-090-843</td>
			<td style="width:401px;">for petroleum jelly syringe</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">3-mL slip tip syringe</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">14-823-436</td>
			<td style="width:401px;">for petroleum jelly syringe</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">20g 3.8 cm slip tip needle</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">14-826-5B</td>
			<td style="width:401px;">for petroleum jelly syringe</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">plain 7 cm X 2.5 cm microscope slide</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">12-550-A3</td>
			<td style="width:401px;">for eyecup dissection, slicing chamber</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">Seche Vite clear nail polish</td>
			<td style="width:239px;">Amazon</td>
			<td style="width:181px;">B00150LT40</td>
			<td style="width:401px;">for slicing chamber</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">18 mm X 18 mm #1 glass coverslips</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td>12-542A</td>
			<td style="width:401px;">for imaging ladders</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">unflavored dental wax</td>
			<td style="width:239px;">Amazon</td>
			<td style="width:181px;">B01K8WNL5A</td>
			<td style="width:401px;">for imaging ladders</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">double edge razor blades</td>
			<td style="width:239px;">Stoelting</td>
			<td style="width:181px;">51427</td>
			<td style="width:401px;">for tissue slicing</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">tissue slicer with digital micrometer</td>
			<td style="width:239px;">Stoelting</td>
			<td style="width:181px;">51415</td>
			<td style="width:401px;">for tissue slicing</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:332px;">filter paper - white gridded mixed cellulose, 13 mm diameter, 0.45 &#181;m pore size</td>
			<td style="width:239px;">EMD Millipore</td>
			<td style="width:181px;">HAWG01300</td>
			<td style="width:401px;">filter paper for mounting retinas</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">10 cm petri dish</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">FB0875712</td>
			<td style="width:401px;">for fish euthanasia, dissection, imaging ladder assembly</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">15 cm plain-tipped wood applicator stick</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">23-400-112</td>
			<td style="width:401px;">for wire eye loop tool</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">30g (0.25 mm diameter) tungsten wire</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">AA10408G6</td>
			<td style="width:401px;">for wire eye loop tool</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">D-glucose</td>
			<td style="width:239px;">Sigma Aldrich</td>
			<td style="width:181px;">G8270</td>
			<td style="width:401px;">component of supplement stock solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">sodium L-lactate</td>
			<td style="width:239px;">Sigma Aldrich</td>
			<td style="width:181px;">L7022</td>
			<td style="width:401px;">component of supplement stock solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">sodium pyruvate</td>
			<td style="width:239px;">Sigma Aldrich</td>
			<td style="width:181px;">P2256</td>
			<td style="width:401px;">component of supplement stock solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">L-glutamine</td>
			<td style="width:239px;">Sigma Aldrich</td>
			<td style="width:181px;">G3126</td>
			<td style="width:401px;">component of supplement stock solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">&#160;L-glutathione, reduced</td>
			<td style="width:239px;">Sigma Aldrich</td>
			<td style="width:181px;">G4251</td>
			<td style="width:401px;">component of supplement stock solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">L-ascorbic acid</td>
			<td style="width:239px;">Sigma Aldrich</td>
			<td style="width:181px;">A5960</td>
			<td style="width:401px;">component of supplement stock solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">NaCl</td>
			<td style="width:239px;">Sigma Aldrich</td>
			<td style="width:181px;">S7653</td>
			<td style="width:401px;">component of Ringer&#39;s solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">KCl</td>
			<td style="width:239px;">Sigma Aldrich</td>
			<td style="width:181px;">P9333</td>
			<td style="width:401px;">component of Ringer&#39;s solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">CaCl2 &#183; 2H2O</td>
			<td style="width:239px;">Sigma Aldrich</td>
			<td style="width:181px;">C3881</td>
			<td style="width:401px;">component of Ringer&#39;s solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">NaH2PO4</td>
			<td style="width:239px;">Sigma Aldrich</td>
			<td style="width:181px;">S8282</td>
			<td style="width:401px;">component of Ringer&#39;s solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">MgCl2 &#183; 6H2O</td>
			<td style="width:239px;">Sigma Aldrich</td>
			<td style="width:181px;">M0250</td>
			<td style="width:401px;">component of Ringer&#39;s solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">HEPES</td>
			<td style="width:239px;">Sigma Aldrich</td>
			<td style="width:181px;">H3375</td>
			<td style="width:401px;">component of Ringer&#39;s solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Tris base</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">BP152</td>
			<td style="width:401px;">component of Na+-free Ringer&#39;s solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">6 N HCl</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">02-003-063</td>
			<td style="width:401px;">component of Na+-free Ringer&#39;s solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">KH2PO4</td>
			<td style="width:239px;">Sigma Aldrich</td>
			<td style="width:181px;">P5655</td>
			<td style="width:401px;">component of Na+-free Ringer&#39;s solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">50 mL conical centrifuge tube</td>
			<td style="width:239px;">Denville Scientific</td>
			<td style="width:181px;">C1062-P</td>
			<td style="width:401px;">container for Ringer&#39;s solution</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">Vannas scissors - 8 cm, angled 5 mm blades</td>
			<td style="width:239px;">World Precision Instruments</td>
			<td style="width:181px;">501790</td>
			<td style="width:401px;">micro-scissors for eyecup dissection</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:332px;">Swiss tweezers - #5, 11 cm, straight, 0.06 X 0.07 mm tips</td>
			<td style="width:239px;">World Precision Instruments</td>
			<td style="width:181px;">504510</td>
			<td style="width:401px;">fine forceps for eyecup dissection and slice manipulation</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">single edge razor blades</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">12-640</td>
			<td style="width:401px;">for eyecup dissection and trimming filter paper</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">EMD Millipore filter forceps</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">XX6200006P</td>
			<td style="width:401px;">flat forceps for handling wet filter paper</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:332px;">C12 558/568 BODIPY</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">D3835</td>
			<td style="width:401px;">stains live cell nuclei; incubate 5 &#181;g/mL for 15 min at room temperature</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:332px;">propidium iodide (PI)</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">P3566</td>
			<td style="width:401px;">stains dead cell nuclei; incubate 5 &#181;g/mL for 20 min at room temperature</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:332px;">Hoechst 33342</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">62249</td>
			<td style="width:401px;">stains live cell nuclei; incubate 5 &#181;g/mL for 20 min at room temperature</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:332px;">Tetramethylrhodamine, methyl ester (TMRM)</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">T668</td>
			<td style="width:401px;">stains functional, negatively-charged mitochondria; incubate 1 nM for 30 min at room temperature</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">tissue perfusion chamber</td>
			<td style="width:239px;">Cell MicroControls</td>
			<td style="width:181px;">BT-1-18/BT-1-18BV [-SY]</td>
			<td style="width:401px;">imaging chamber for injection or perfusion</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:43px;width:332px;">2-(N-(7-Nitrobenz-2-oxa-1,3-diazol-4-yl)Amino)-2-Deoxyglucose (NBDG)</td>
			<td style="width:239px;">Fisher Scientific</td>
			<td style="width:181px;">N13195</td>
			<td style="width:401px;">fluorescent glucose analog adminitered orally to zebrafish 30 min prior to euthanasia</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:332px;">Olympus laser scanning confocal microscope</td>
			<td style="width:239px;">Olympus</td>
			<td style="width:181px;">FV1000</td>
			<td style="width:401px;">confocal microscope for visualizing fluorescence of slices at single-cell resolution</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:332px;">Carbonyl cyanide 3-chlorophenylhydrazone (CCCP)</td>
			<td style="width:239px;">Sigma Aldrich</td>
			<td style="width:181px;">C2759</td>
			<td style="width:401px;">experimental reagent which ablates mitochondrial respiration; treat slices to a final concentration of 1 &#181;M</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:332px;">miniature aspirator positioner</td>
			<td style="width:239px;">Cell MicroControls</td>
			<td style="width:181px;">FL-1</td>
			<td style="width:401px;">for perfusion</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:332px;">perfusion manifold, gas bubbler manifold, flow valve, 60cc syringe holder</td>
			<td style="width:239px;">Warner Instruments</td>
			<td style="width:181px;">various</td>
			<td style="width:401px;">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

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, ...

Ex Vivo Imaging of Postnatal Cerebellar Granule Cell Migration Using Confocal Macroscopy

During postnatal development, immature granule cells (excitatory interneurons) exhibit tangential migration in the external granular layer, and then radial migration in the molecular layer and the Purkinje cell layer to reach the internal granular layer of the cerebellar cortex. Default in migratory processes induces either cell death or misplacement of the neurons, leading to deficits in diverse cerebellar functions. Centripetal granule cell migration involves several mechanisms, such as chemotaxis and extracellular matrix degradation, to guide the cells towards their final position, but the factors that regulate cell migration in each cortical layer are only partially known. In our method, acute cerebellar slices are prepared from P10 rats, granule cells are labeled with a fluorescent cytoplasmic marker and tissues are cultured on membrane inserts from 4 to 10 hr before starting real-time monitoring of cell migration by confocal macroscopy at 37 &#176;C in the presence of CO2. During their migration in the different cortical layers of the cerebellum, granule cells can be exposed to neuropeptide agonists or antagonists, protease inhibitors, blockers of intracellular effectors or even toxic substances such as alcohol or methylmercury to investigate their possible role in the regulation of neuronal migration.

Ex Vivo Imaging of Postnatal Cerebellar Granule Cell Migration Using Confocal Macroscopy

During postnatal cerebellum development, immature granule cells originating from the germinal zone exhibit distinct modalities of migration to reach their final destination and to establish neuronal networks. This protocol describes the preparation of cerebellar slices and the confocal macroscopic approach used to investigate the factors that regulate neuronal migration.

During postnatal development, immature granule cells (excitatory interneurons) exhibit tangential migration in the external granular layer, and then ...

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated proteins and subsequent manipulation of neural activity by light. Channelrhodopsins (ChRs) are light-gated cation-channels and when fused to a fluorescent protein their expression allows for visualization and concurrent activation of specific cell types and their axonal projections in defined areas of the brain. Via stereotactic injection of viral vectors, ChR fusion proteins can be constitutively or conditionally expressed in specific cells of a defined brain region, and their axonal projections can subsequently be studied anatomically and functionally via ex vivo optogenetic activation in brain slices. This is of particular importance when aiming to understand synaptic properties of connections that could not be addressed with conventional electrical stimulation approaches, or in identifying novel afferent and efferent connectivity that was previously poorly understood. Here, a few examples illustrate how this technique can be applied to investigate these questions to elucidating fear-related circuits in the amygdala. The amygdala is a key region for acquisition and expression of fear, and storage of fear and emotional memories. Many lines of evidence suggest that the medial prefrontal cortex (mPFC) participates in different aspects of fear acquisition and extinction, but its precise connectivity with the amygdala is just starting to be understood. First, it is shown how ex vivo optogenetic activation can be used to study aspects of synaptic communication between mPFC afferents and target cells in the basolateral amygdala (BLA). Furthermore, it is illustrated how this ex vivo optogenetic approach can be applied to assess novel connectivity patterns using a group of GABAergic neurons in the amygdala, the paracapsular intercalated cell cluster (mpITC), as an example.

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are widely used to manipulate neural activity and assess the consequences for brain function. Here, a technique is outlined that&#160;upon in vivo expression of the optical activator Channelrhodopsin, allows for ex vivo analysis of synaptic properties of specific long range and local neural connections in fear-related circuits.

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated ...

Optical Coherence Tomography: Imaging Mouse Retinal Ganglion Cells In Vivo

Structural changes in the retina are common manifestations of ophthalmic diseases. Optical coherence tomography (OCT) enables their identification in vivo&#8212;rapidly, repetitively, and at a high resolution. This protocol describes OCT imaging in the mouse retina as a powerful tool to study optic neuropathies (OPN). The OCT system is an interferometry-based, non-invasive alternative to common post mortem histological assays. It provides a fast and accurate assessment of retinal thickness, allowing the possibility to track changes, such as retinal thinning or thickening. We present the imaging process and analysis with the example of the Opa1delTTAG mouse line. Three types of scans are proposed, with two quantification methods: standard and homemade calipers. The latter is best for use on the peripapillary retina during radial scans; being more precise, is preferable for analyzing thinner structures. All approaches described here are designed for retinal ganglion cells (RGC) but are easily adaptable to other cell populations. In conclusion, OCT is efficient in mouse model phenotyping and has the potential to be used for the reliable evaluation of therapeutic interventions.

Optical Coherence Tomography: Imaging Mouse Retinal Ganglion Cells In Vivo

This manuscript describes a protocol for the in vivo imaging of the mouse retina with high-resolution spectral domain optical coherence tomography (SD-OCT). It focuses on retinal ganglion cells (RGC) in the peripapillary region, with several scanning and quantifying approaches described.

Structural changes in the retina are common manifestations of ophthalmic diseases. Optical coherence tomography (OCT) enables their identification in ...

Assessment of Long-term Depression Induction in Adult Cerebellar Slices

Synaptic plasticity provides a mechanism for learning and memory. For cerebellar motor learning, long-term depression (LTD) of synaptic transmissions from parallel fibers (PF) to Purkinje cells (PC) is considered the basis for motor learning, and deficiencies of both LTD and motor learning are observed in various gene-manipulated animals. Common motor learning sets, such as adaptation of the optokinetic reflex (OKR), the vestibular-ocular reflex (VOR), and rotarod test were used for evaluation of motor learning ability. However, results obtained from the GluA2-carboxy terminus modified knock-in mice demonstrated normal adaptation of the VOR and the OKR, despite lacking PF-LTD. In that report, induction of LTD was only attempted using one type of stimulation protocol at room temperature. Thus, conditions to induce cerebellar LTD were explored in the same knock-in mutants using various protocols at near physiological temperature. Finally, we found stimulation protocols, by which LTD could be induced in these gene-manipulated mice. In this study, a set of protocols are proposed to evaluate LTD-induction, which will more accurately allow examination of the causal relationship between LTD and motor learning. In conclusion, experimental conditions are crucial when evaluating LTD in gene-manipulated mice.

In some gene-manipulated animals, using a single protocol may fail to induce LTD in cerebellar Purkinje cells, and there may be a discrepancy between LTD and motor learning. Multiple protocols are necessary to assess LTD-induction in gene-manipulated animals. Standard protocols are shown.

Synaptic plasticity provides a mechanism for learning and memory. For cerebellar motor learning, long-term depression (LTD) of synaptic transmissions from ...

Ex Vivo Oculomotor Slice Culture from Embryonic GFP-Expressing Mice for Time-Lapse Imaging of Oculomotor Nerve Outgrowth

Accurate eye movements are crucial for vision, but the development of the ocular motor system, especially the molecular pathways controlling axon guidance, has not been fully elucidated. This is partly due to technical limitations of traditional axon guidance assays. To identify additional axon guidance cues influencing the oculomotor nerve, an ex vivo slice assay to image the oculomotor nerve in real-time as it grows towards the eye was developed. E10.5 IslMN-GFP embryos are used to generate ex vivo slices by embedding them in agarose, slicing on a vibratome, then growing them in a microscope stage-top incubator with time-lapse photomicroscopy for 24-72 h. Control slices recapitulate the in vivo timing of outgrowth of axons from the nucleus to the orbit. Small molecule inhibitors or recombinant proteins can be added to the culture media to assess the role of different axon guidance pathways. This method has the advantages of maintaining more of the local microenvironment through which axons traverse, not axotomizing the growing axons, and assessing the axons at multiple points along their trajectory. It can also identify effects on specific subsets of axons. For example, inhibition of CXCR4 causes axons still within the midbrain to grow dorsally rather than ventrally, but axons that have already exited ventrally are not affected.

An ex vivo slice assay allows oculomotor nerve outgrowth to be imaged in real time. Slices are generated by embedding E10.5 IslMN:GFP embryos in agarose, slicing on a vibratome, and growing in a stage-top incubator. The role of axon guidance pathways is assessed by adding inhibitors to the culture media.

Accurate eye movements are crucial for vision, but the development of the ocular motor system, especially the molecular pathways controlling axon ...

Minimizing Hypoxia in Hippocampal Slices from Adult and Aging Mice

Acute hippocampal slices have enabled generations of neuroscientists to explore synaptic, neuronal, and circuit properties in detail and with high fidelity. Exploration of LTP and LTD mechanisms, single neuron dendritic computation, and experience-dependent changes in circuitry, would not have been possible without this classical preparation. However, with a few exceptions, most basic research using acute hippocampal slices has been performed using slices from rodents of relatively young ages, ~P20-P40, even though synaptic and intrinsic excitability mechanisms have a long developmental tail that reaches past P60. The main appeal of using young hippocampal slices is preservation of neuronal health aided by higher tolerance to hypoxic damage. However, there is a need to understand neuronal function at more mature stages of development, further accentuated by the development of various animal models of neurodegenerative diseases that require an aging brain preparation. Here we describe a modification to an acute hippocampal slice preparation that reliably delivers healthy slices from adult and aging mouse hippocampi. The protocol&#8217;s critical steps are transcardial perfusion and cutting with ice-cold sodium-free NMDG-aSCF. Together, these steps attenuate the hypoxia-induced drop in ATP upon decapitation, as well as cytotoxic edema caused by passive sodium fluxes. We demonstrate how to cut transversal slices of hippocampus plus cortex using a vibrating microtome. Acute hippocampal slices obtained in this way are reliably healthy over many hours of recording, and are appropriate for both field-recordings and targeted patch-clamp recordings, including targeting of fluorescently labeled neurons.

This is a protocol for acute slice preparation from adult and aging mouse hippocampi that takes advantage of transcardial perfusion and slice cutting with ice-cold NMDG-aCSF to reduce hypoxic damage to the tissue. The resulting slices stay healthy over many hours, and are suitable for long-term patch-clamp and field-recordings.

Acute hippocampal slices have enabled generations of neuroscientists to explore synaptic, neuronal, and circuit properties in detail and with high ...

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are optimized to minimize their impact on behavior. This is first achieved by using a holder that minimizes movement hindrances. The back of the fly&#8217;s head is glued to this holder at an angle that allows optical access to the whole brain while retaining the fly&#8217;s ability to walk, groom, smell, taste and see. The back of the head is dissected to remove tissues in the optical path and muscles responsible for head movement artefacts. The fly brain can subsequently be imaged to record brain activity, for instance using calcium or voltage indicators, during specific behaviors such as walking or grooming, and in response to different stimuli. Once the challenging dissection, which requires considerable practice, has been mastered, this technique allows to record rich data sets relating whole brain activity to behavior and stimulus responses.

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

We present a method specifically tailored to image the whole brain of adult Drosophila during behavior and in response to stimuli. The head is positioned to allow optical access to the whole brain, while the fly can move its legs and the antennae, the tip of the proboscis, and the eyes can receive sensory stimuli.

We present a method developed specifically to image the whole Drosophila brain during ongoing behavior such as walking. Head fixation and dissection are ...

Isolation of Adult Human Astrocyte Populations from Fresh-Frozen Cortex Using Fluorescence-Activated Nuclei Sorting

The complexity of human astrocytes remains poorly defined in primary human tissue, requiring better tools for their isolation and molecular characterization. Fluorescence-activated nuclei sorting (FANS) can be used to successfully isolate and study human neuronal nuclei (NeuN+) populations from frozen archival tissue, thereby avoiding problems associated with handling&#160;fresh tissue. However, efforts to similarly isolate astroglia from the non-neuronal (NeuN-) element are lacking. A recently developed and validated immunotagging strategy uses three transcription factor antibodies to simultaneously isolate enriched neuronal (NeuN+), astrocyte (paired box protein 6 (PAX6)+NeuN-), and oligodendrocyte progenitor (OLIG2+NeuN-) nuclei populations from non-diseased, fresh (unfixed) snap-frozen&#160;postmortem human temporal neocortex tissue.
This technique was shown to be useful for the characterization of cell type-specific transcriptome alterations in primary pathological epilepsy neocortex. Transcriptomic analyses confirmed that PAX6+NeuN- sorted populations are robustly enriched for pan-astrocyte markers and capture astrocytes in both resting and reactive conditions. This paper describes the FANS methodology for the isolation of astrocyte-enriched nuclei populations from fresh-frozen human cortex, including tissue dissociation into single-nucleus (sn) suspension; immunotagging of nuclei with anti-NeuN and anti-PAX6 fluorescently conjugated antibodies; FANS gating strategies and quality control metrics for optimizing sensitivity and specificity during sorting and for confirming astrocyte enrichment; and recommended procurement for downstream transcriptome and chromatin accessibility sequencing at bulk or sn resolution. This protocol is applicable for non-necrotic, fresh-frozen, human cortical specimens with various pathologies and recommended postmortem tissue collection within 24 h.

We have developed a method that enriches for and isolates human astrocyte populations from fresh-frozen tissue for use in downstream molecular analyses.

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

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Assessment of Long-term Depression Induction in Adult Cerebellar Slices

Ex Vivo Oculomotor Slice Culture from Embryonic GFP-Expressing Mice for Time-Lapse Imaging of Oculomotor Nerve Outgrowth

Minimizing Hypoxia in Hippocampal Slices from Adult and Aging Mice

Preparing Adult Drosophila melanogaster for Whole Brain Imaging during Behavior and Stimuli Responses

Isolation of Adult Human Astrocyte Populations from Fresh-Frozen Cortex Using Fluorescence-Activated Nuclei Sorting