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Method Article

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

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DOI:

10.3791/56977

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May 9th, 2018

 ,  ,  , 
1Department of Biochemistry, University of Washington, 2Department of Ophthalmology, University of Washington

In This Article

Summary

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.

Abstract

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'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 Müller 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.

Introduction

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 Müller 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.

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Protocol

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 800x800 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.

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Results

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 ...

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Discussion

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...

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Disclosures

The authors declare that they have no competing financial interests.

Acknowledgements

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.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
zebrafishUniveristy of Washington South Lake Union Aquatics Facilitystocks maintained in-house as stable transgenic lines
petroleum jellyFisher Scientific19-090-843for petroleum jelly syringe
3-mL slip tip syringeFisher Scientific14-823-436for petroleum jelly syringe
20g 3.8 cm slip tip needleFisher Scientific14-826-5Bfor petroleum jelly syringe
plain 7 cm X 2.5 cm microscope slideFisher Scientific12-550-A3for eyecup dissection, slicing chamber
Seche Vite clear nail polishAmazonB00150LT40for slicing chamber
18 mm X 18 mm #1 glass coverslipsFisher Scientific12-542Afor imaging ladders
unflavored dental waxAmazonB01K8WNL5Afor imaging ladders
double edge razor bladesStoelting51427for tissue slicing
tissue slicer with digital micrometerStoelting51415for tissue slicing
filter paper - white gridded mixed cellulose, 13 mm diameter, 0.45 µm pore sizeEMD MilliporeHAWG01300filter paper for mounting retinas
10 cm petri dishFisher ScientificFB0875712for fish euthanasia, dissection, imaging ladder assembly
15 cm plain-tipped wood applicator stickFisher Scientific23-400-112for wire eye loop tool
30g (0.25 mm diameter) tungsten wireFisher ScientificAA10408G6for wire eye loop tool
D-glucoseSigma AldrichG8270component of supplement stock solution
sodium L-lactateSigma AldrichL7022component of supplement stock solution
sodium pyruvateSigma AldrichP2256component of supplement stock solution
L-glutamineSigma AldrichG3126component of supplement stock solution
 L-glutathione, reducedSigma AldrichG4251component of supplement stock solution
L-ascorbic acidSigma AldrichA5960component of supplement stock solution
NaClSigma AldrichS7653component of Ringer's solution
KClSigma AldrichP9333component of Ringer's solution
CaCl2 · 2H2OSigma AldrichC3881component of Ringer's solution
NaH2PO4Sigma AldrichS8282component of Ringer's solution
MgCl2 · 6H2OSigma AldrichM0250component of Ringer's solution
HEPESSigma AldrichH3375component of Ringer's solution
Tris baseFisher ScientificBP152component of Na+-free Ringer's solution
6 N HClFisher Scientific02-003-063component of Na+-free Ringer's solution
KH2PO4Sigma AldrichP5655component of Na+-free Ringer's solution
50 mL conical centrifuge tubeDenville ScientificC1062-Pcontainer for Ringer's solution
Vannas scissors - 8 cm, angled 5 mm bladesWorld Precision Instruments501790micro-scissors for eyecup dissection
Swiss tweezers - #5, 11 cm, straight, 0.06 X 0.07 mm tipsWorld Precision Instruments504510fine forceps for eyecup dissection and slice manipulation
single edge razor bladesFisher Scientific12-640for eyecup dissection and trimming filter paper
EMD Millipore filter forcepsFisher ScientificXX6200006Pflat forceps for handling wet filter paper
C12 558/568 BODIPYFisher ScientificD3835stains live cell nuclei; incubate 5 µg/mL for 15 min at room temperature
propidium iodide (PI)Fisher ScientificP3566stains dead cell nuclei; incubate 5 µg/mL for 20 min at room temperature
Hoechst 33342Fisher Scientific62249stains live cell nuclei; incubate 5 µg/mL for 20 min at room temperature
Tetramethylrhodamine, methyl ester (TMRM)Fisher ScientificT668stains functional, negatively-charged mitochondria; incubate 1 nM for 30 min at room temperature
tissue perfusion chamberCell MicroControlsBT-1-18/BT-1-18BV [-SY]imaging chamber for injection or perfusion
2-(N-(7-Nitrobenz-2-oxa-1,3-diazol-4-yl)Amino)-2-Deoxyglucose (NBDG)Fisher ScientificN13195fluorescent glucose analog adminitered orally to zebrafish 30 min prior to euthanasia
Olympus laser scanning confocal microscopeOlympusFV1000confocal microscope for visualizing fluorescence of slices at single-cell resolution
Carbonyl cyanide 3-chlorophenylhydrazone (CCCP)Sigma AldrichC2759experimental reagent which ablates mitochondrial respiration; treat slices to a final concentration of 1 µM
miniature aspirator positionerCell MicroControlsFL-1for perfusion
perfusion manifold, gas bubbler manifold, flow valve, 60cc syringe holderWarner Instrumentsvariousfor perfusion

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