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

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.

1. 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.
   1. 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.
2. 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.
3. Make a reusable slicing chamber on a plain 7 cm X 2.5 cm microscope slide.
   1. 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.
4. Prepare imaging ladders on cover slips to hold slices during imaging. The slices will form the "rungs" of the ladder.
   1. 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.
5. Ready the tissue slicer.
   1. 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.
   2. 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.
   3. 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 ¼ 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.
6. 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.
7. 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.
8. 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.
9. Prepare Ringer's solution.
   1. Thaw 50X supplement stock solution (Table 1) and add it fresh to HEPES-buffered, non-bicarbonate Ringer's solution (Table 2) the morning of the experiment; dilute 200 µL of supplement stock solution in every 10 mL of Ringer's solution in a conical centrifuge tube or sterile glass bottle. For static imaging experiments, prepare at least 30 mL of Ringer's solution per experiment. The volume of Ringer's solution needed for perfusion experiments will depend on the flow rate and total experiment time.
   2. 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.
   3. Oxygenate supplemented Ringer'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'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.
   4. If other solutions are being used in the experiment, such as Na⁺-free Ringer's solution (Table 3), repeat steps 1.9.1.-1.9.3.
10. Gather Petri dishes, forceps, micro-scissors and other tools near the dissection microscope, and prepare a fish water ice bath for zebrafish euthanasia.

2. Preparing retinal slices (see Figure 1)

1. 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.
2. 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.
3. Transfer the eye using forceps to a Petri dish of cold Ringer'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.
4. Dissect the eyecups under a low power dissection microscope in a drop of cold Ringer's solution on a plain glass slide in a Petri dish.
   1. 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.
   2. 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.
   3. 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.
5. Flat mount retina on filter paper.
   1. Wet a piece of filter paper with Ringer'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's solution to cover both the filter paper and tissue.
   2. 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.
   3. 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'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.
6. 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'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.
7. 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'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.
8. 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'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 µg/mL for 15 min at room temperature (typically 23-27 °C), followed by a wash in excess Ringer's solution.
9. 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 µm slices.
10. Assemble the imaging ladder.
    1. 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's solution to submerge its contents.
    2. 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° and bury the filter paper edges in petroleum jelly.
    3. 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).
11. If desired, load dyes into retinal slices at this stage and wash in excess Ringer'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 µg/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.
12. While the slices are staining, prepare the imaging chamber and injection apparatus. Use the syringe to flush the tubing with Ringer'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.
13. 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's solution to cover slices.

3. Imaging retinal slices

1. 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.
2. 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.
3. 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 800×800 pixel resolution, 2 µs/pixel scan speed, and a 10-s frame rate for imaging calcium dynamics in photoreceptors with GCaMP3 and a red dye.
   1. 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.
4. 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 µM 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.
5. 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 < 5 µm.

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.

1. Instead of petroleum jelly ladders described in step 1.4, use sturdier wax ladders to hold retinal slices steady during solution flow.
   1. 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.
   2. 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).
2. 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's solution, ensure all lines flow when opened and purge large bubbles.
3. 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.
4. 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'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's solution flowing while selecting slices in step 3.4.
5. 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.
   1. For instance, to deplete extracellular Na⁺ from the imaging chamber, use two syringe reservoirs filled with Ringer's solution or Na⁺-free solution (Table 3). Flow Ringer'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).
6. 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.