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

Summary

The article explains how to surgically remove eyes from living zebrafish larvae as the first step toward investigating how retinal input influences optic tectum growth and development. In addition, the article provides information about larval anesthetization, fixation, and brain dissection, followed by immunohistochemistry and confocal imaging.

Abstract

Zebrafish exhibit remarkable life-long growth and regenerative abilities. For example, specialized stem cell niches established during embryogenesis support continuous growth of the entire visual system, both in the eye and the brain. Coordinated growth between the retinae and the optic tectum ensures accurate retinotopic mapping as new neurons are added in the eyes and brain. To address whether retinal axons provide crucial information for regulating tectal stem and progenitor cell behaviors such as survival, proliferation, and/or differentiation, it is necessary to be able to compare innervated and denervated tectal lobes within the same animal and across animals.

Surgical removal of one eye from living larval zebrafish followed by observation of the optic tectum achieves this goal. The accompanying video demonstrates how to anesthetize larvae, electrolytically sharpen tungsten needles, and use them to remove one eye. It next shows how to dissect brains from fixed zebrafish larvae. Finally, the video provides an overview of the protocol for immunohistochemistry and a demonstration of how to mount stained embryos in low-melting-point agarose for microscopy.

Introduction

The goal of this method is to investigate how retinal input influences the growth and development of the optic tectum, the visual processing center in the zebrafish brain. By removing one eye and then comparing the two sides of the optic tectum, tectal changes within the same specimen can be observed and normalized, enabling comparison across multiple specimens. Modern molecular approaches combined with this technique will yield insights into the mechanisms underlying visual system growth and development, as well as axonal degeneration and regeneration.

Sensory systems - visual, auditory, and somatosensory - gather information from external organs and relay that information to the central nervous system, generating "maps" of the external world across the midbrain^1,2. Vision is the dominant sensory modality for nearly all vertebrates, including many fishes. The retina, the neural tissue in the eye, gathers information with a neuronal circuit consisting primarily of photoreceptors, bipolar cells, and retinal ganglion cells (RGCs), the projection neurons of the retina. RGCs have long axons that find their way across the inner surface of the retina to the optic nerve head, where they fasciculate and travel together through the brain, ultimately terminating in the visual processing center in the dorsal midbrain. This structure is called the optic tectum in fish and other non-mammalian vertebrates and is homologous to the superior colliculus in mammals³.

The optic tectum is a bilaterally symmetric multilayered structure in the dorsal midbrain. In zebrafish and most other fishes, each lobe of the optic tectum receives visual input solely from the contralateral eye, such that the left optic nerve terminates in the right tectal lobe and the right optic nerve terminates in the left tectal lobe⁴ (Figure 1). Like its mammalian counterpart, the superior colliculus, the optic tectum integrates visual information with other sensory inputs, including audition and somatosensation, controlling shifts in visual attention and eye movements such as saccades^1,5,6. However, unlike the mammalian superior colliculus, the optic tectum continuously generates new neurons and glia from a specialized stem cell niche near the medial and caudal edges of the tectal lobes called the tectal proliferation zone⁷. Maintenance of proliferative progenitors in the optic tectum and other regions of the central nervous system contributes, in part, to the remarkable regenerative capacity documented in zebrafish⁸.

Previous work examining the brains of blind or one-eyed fishes revealed that optic tectum size is directly proportional to the amount of retinal innervation it receives^9,10,11. In adult cave fish, whose eyes degenerate in early embryogenesis, the optic tectum is noticeably smaller than that of closely related, sighted surface fish⁹. Cave fish eye degeneration can be blocked by replacing the endogenous lens with a lens from a surface fish during embryogenesis. When these one-eyed cave fish are reared to adulthood, the innervated tectal lobe contains approximately 10% more cells than the non-innervated tectal lobe⁹. Similarly, in larval killifish that were incubated with chemical treatments to generate eyes of different sizes within the same individual, the side of the tectum with more innervation was larger and contained more neurons¹⁰. Evidence from optic nerve crush experiments in adult goldfish indicates that innervation promotes proliferation, with tectal cell proliferation decreasing when innervation was disrupted¹¹.

Confirming and extending these classical studies, several recent reports provide data suggesting that proliferation in response to innervation is modulated, at least in part, by the BDNF-TrkB pathway^12,13. Many open questions about optic tectum growth and development remain, including how a developing sensory system copes with injury and axon degeneration, which cellular and molecular signals enable retinal input to regulate optic tectum growth, when these mechanisms become active, and whether innervation-linked proliferation and differentiation enable the retina and its target tissue to coordinate growth rates and ensure accurate retinotopic mapping. In addition, there are much larger questions about activity-dependent development that can be addressed by interrogating the zebrafish visual system with surgical approaches such as the one described below.

To investigate the cellular and molecular mechanisms by which neural activity, specifically from visual input, alters cell survival and proliferation, the described approach directly compares innervated and denervated tectal lobes (Figure 1) within individual zebrafish larvae. This method allows for the documentation of RGC axon degeneration in the optic tectum and confirmation that the number of mitotic cells correlates with innervation.

Figure 1: Sketches of zebrafish larvae before and after unilateral eye removal. (A) Drawing of 5 dpf larvae as viewed under a dissecting microscope. Each larva is embedded in low-melting-point agarose and oriented laterally before a tungsten needle with a sharp, hooked tip is used to scoop out the eye facing up (left eye in this example). (B) Drawing of the dorsal view of a 9 dpf larva resulting from the surgery depicted in A. Only three highly schematized RGC axons from the right eye are shown defasciculating and connecting with neurons in the left tectal lobe. Abbreviations: dpf = days post fertilization; dps = days post surgery; RGC = retinal ganglion cells. Please click here to view a larger version of this figure.

Protocol

The methods in this paper were conducted in accordance with guidelines and approval of the Institutional Animal Care and Use Committees of Reed College and University College London. See the Table of Materials for details about zebrafish strains used in this study.

1. Prepare materials and tools

1. Make solutions.
   1. Make embryo medium (E3¹⁴) by diluting a 60x stock (300 mM NaCl, 10.2 mM KCl, 20 mM CaCl₂-dihydrate, and 20 mM MgCl₂-hexahydrate in deionized water, autoclaved, and stored at room temperature). Add 160 mL of 60x E3 to 10 L of deionized water. Store at room temperature.
   2. Make 1% low melting point (LMP) agarose in E3. Dissolve 1 g of LMP agarose into 100 mL of E3 by boiling in the microwave for 1-2 min on high power. Aliquot molten agarose into 1.7 mL microfuge tubes, and store at 4 °C. Boil in a water bath and hold in a 40 °C heat block when ready to use for embedding.
   3. Make Marc's Modified Ringers (MMR¹⁵) isotonic solution by diluting a 10x stock (1 M NaCl, 20 mM KCl, 10 mM MgSO₄, 20 mM CaCl₂-dihydrate, 50 mM HEPES, pH adjusted to 7.5 with 10 M NaOH, then autoclaved, and stored at room temperature). Add 10 mL of 10x MMR to 90 mL of deionized water.
2. Pour Sylgard plates according to the manufacturer's instructions, allowing them to set and dry for several days¹⁶.
3. Electrolytically sharpen tungsten needles (protocol modified from ¹⁷).
   1. First, prepare a 10% (w/v) KOH solution by adding 200 mL of deionized water to a wide-mouthed glass jar with a screw-top lid. Slowly stir in 20 g of KOH pellets.
      NOTE: This caustic solution is highly exothermic. Wear gloves and eye protection, and stir the solution in the hood.
   2. Cut a 2-3 cm length of tungsten wire and insert it into the needle holder.
   3. Connect the cathode and anode wires to the power supply. Attach the alligator clip at the end of the cathode wire to a partially straightened paper clip and insert the paper clip into the KOH solution, attaching it to the side of the jar.
   4. Next, attach the alligator clip of the anode wire to the neck of the needle holder. Turn the power on (to ~20 V) and dip the tungsten wire into the KOH solution, pulling the wire out of the solution at an angle to electrolytically sharpen the wire into a fine tip.
      NOTE: Bubbles will come from the paper clip as the reaction proceeds.
   5. Check the tip of the needle under the dissecting microscope to ensure that it is sharp enough.
   6. Rinse the needle with deionized water to remove all KOH residue. Make several needles ahead of time and keep them until the larval surgeries and dissections.
4. Make chambered slides for imaging zebrafish specimens with an upright confocal microscope.
   1. Obtain glass microscope slides and two-part epoxy resin (see the Table of Materials). Mix the epoxy according to the instructions on the package, and paint thick rings or rectangles on the glass slides. Allow the slides to cure and dry in the hood for at least 24 h before use.

2. Embryo collection and rearing

1. Pair adult male and female fish of the desired genotypes in breeding boxes in the late afternoon/early evening. The next morning, after they have spawned, collect newly fertilized eggs with a mesh strainer and transfer them to E3 in 100 mm Petri dishes.
2. Incubate the embryos at ~27.5 °C with a 14 h light/10 h dark cycle until the day of the surgery. Change E3 daily or as needed.
3. Feed the larvae using a dropper full of harvested rotifers once a day, beginning either at 5 days post fertilization (dpf) or one day after surgery. After the larvae have several hours to eat, change the E3 to remove any remaining rotifers and waste products.
   ​NOTE: Do not feed larvae on the day of surgery.

3. Prepare larvae for surgery

1. On the day of surgery, use a wide-bore, glass Pasteur pipette to transfer 10-15 larvae to a 35 mm Petri dish filled with fresh E3.
2. Anesthetize the larvae by adding 3-5 drops of 0.4% w/v Tricaine solution (0.4 g Tricaine dissolved in 210 mM Tris, pH 9; final solution adjusted to a pH of 7.4 if necessary¹⁸) to the dish for a final concentration of ~0.0015% w/v Tricaine. Look for lack of response to touch to determine if the larvae are adequately anesthetized. If they are still responsive to touch after ~3 min, add 2-3 more drops of 0.4% w/v Tricaine solution and reassess.
   NOTE: At this stage of development, it is possible to monitor blood flow and heart rate in addition to motility under the dissecting microscope.
3. Immobilize the anesthetized zebrafish larvae by embedding them in 1% LMP agarose dissolved in E3.
   1. First, place the lid of a 35 mm Petri dish face up under the dissecting microscope. Next, take one larva up into a narrow-bore, glass Pasteur pipette with only a small amount of E3.
   2. Then, aspirate ~200 µL of melted, warm (~40 °C) 1% LMP agarose into the pipette with the larva. Finally, squirt the larva and agarose onto the upside-down Petri dish lid.
   3. Use a dull tungsten needle to quickly but gently maneuver the larva so that it is lateral, with one eye facing up. Wait for the agarose to set (~5 min).
      ​NOTE: If the agarose hardens while the larva is not lateral, free it from the agarose (as described in step 5) and return it to the dish with E3 and tricaine.

4. Eye-removal

1. Once the agarose is set, and the larva is positioned laterally, use the tip of a very fine, electrolytically sharpened tungsten needle to pierce the skin around the eye, following the edge of the eye orbit.
2. Next, slide the edge of the needle (not the tip) under the eye from the temporal-ventral side of the eye. Use controlled pressure to release the eye from the socket.
3. Use very fine surgical forceps to remove the eye by pinching the optic nerve and pushing the eye from medial to lateral. Alternatively, simply keep pressing the eye dorsally and anteriorly with the side of the needle, eventually slicing through the optic nerve and releasing the eye.
   ​NOTE: If tissues other than the eye are accidentally stabbed during the surgery, quickly and humanely kill the larva by rapid decapitation.

5. Postoperative care until experimental endpoint

1. After successful eye removal, cover the agarose with MMR solution.
2. Liberate each larva from the agarose by gently brushing a tungsten needle around their head and then around their body while stabilizing the Petri dish lid with forceps.
3. Transfer the one-eyed larvae to a 35 mm Petri dish that contains MMR supplemented with a 1:100 dilution of a penicillin/streptomycin cocktail. Incubate the embryos at ~27.5 °C with a 14 h light/10 h dark cycle.
   ​NOTE: This isotonic solution of MMR supplemented with antibiotics initially aids larval healing and survival. Each dish can hold up to 15 recovering larvae.
4. The next day, return the larvae to E3. Starting the afternoon following surgery, feed larvae (~6 dpf and older) using a dropper full of harvested rotifers once a day. After the larvae have had several hours to eat, change the E3 to remove any remaining rotifers and waste products.

6. Fixing larvae

1. Once the larvae have reached the desired developmental stage for analysis, anesthetize them with a lethal dose of tricaine (~0.4% final concentration) until their hearts have stopped.
2. Pipette up to 30 larvae per 1.5 mL microfuge tube. Remove excess E3 and then add 0.5-1 mL of fixative solution (4% paraformaldehyde (PFA) and 4% sucrose in phosphate-buffered saline (PBS)) to fix the tissue. Incubate the larvae overnight at 4 °C.
3. Wash the larvae with PBS the next day by removing the fixative and rinsing the larvae 3-4 times with PBS. Store the larvae at 4 °C for up to 7 days before dissection.

7. Dissecting larvae to reveal brains (adapted from ¹⁵)

1. Suspend the fixed larvae in PBS droplets on a Sylgard plate under a dissecting microscope. Secure them laterally by placing two tungsten pins (typically old tungsten needles that are less than 2 cm) through the notochord, with one pin just posterior to the pigmented area covering the aorta-gonad-mesonephros (AGM) region and another in line with the end of the yolk extension.
   NOTE: Position the tungsten pins in as few attempts as possible because the tissue will become more friable with each puncture.
2. Use a very sharp tungsten needle and needle-sharp forceps to expose the brain by removing, in this order, the eye(s), ear, jaw, digestive tract, and dorsal cranial skin.
   1. First, remove the eye using a technique similar to the surgical eye-removal in step 4. Then, unpin the larva and flip it to the opposite side, so the other eye is accessible and repeat. Alternatively, poke the needle through the jaw to the other side and remove the eye without unpinning.
      NOTE: Eyes may be collected at this point if analysis of this tissue is desired (similar to ¹⁹).
   2. Next, use the tungsten needle to scratch from the temporal to the ventral side of the ear. In the same action/motion, bring the needle posterior to the jaw and gently pull anteriorly until the ear and jaw are removed.
   3. Use forceps to pull the ventral organs and any remaining yolk out.
   4. Finally, make a shallow incision in the dorsal cranial skin near the junction between the hindbrain and spinal cord. Lift the skin with the forceps and pull it anteriorly and around the telencephalon. If this proves too difficult, start at the initial incision in the hindbrain and pull the skin first laterally and then ventrally and anteriorly, taking great care not to scratch the lateral edges of the tectum.
      NOTE: Because the midbrain is the object of study, the hindbrain can be cut; however, a deep incision may result in decapitation.
   5. Remove any remaining tissue with forceps. Unpin the larva and transfer to PBS in a 1.5 mL microfuge tube.
      ​NOTE: Leave the tail attached to the brain for better visibility when performing wholemount protocols.

8. Brain dehydration and storage

1. Remove PBS from the brains and add 1 mL of PBS containing 0.1% TritonX-100 (PBST). Incubate for 5-10 min at room temperature.
2. Remove the PBST and replace it with a 50:50 methanol:PBST solution. Incubate for 5-10 min at room temperature.
3. Wash the brains with 100% methanol (MeOH) twice for 5 min each at room temperature.
4. Store the brains in 100% MeOH at -20 °C for at least 16 h before performing immunohistochemistry or other wholemount procedures.
   ​NOTE: Brains can be stored in MeOH for up to 2 years at -20 °C.

9. Immunohistochemistry

NOTE: Established protocols for many useful wholemount techniques in zebrafish can be found on ZFIN²⁰. This manuscript provides examples comparing one-eyed and two-eyed larvae that were immunostained with antibodies that recognize either the red fluorescent protein (RFP), which is expressed in optic nerve axons in the Tg[atoh7:RFP] line (Figure 2), or phosphorylated histone H3 (PH3), which highlights mitotic cells (Figure 3). A standard immunohistochemistry protocol for wholemount embryos and larvae is summarized below.

1. First, rehydrate the larval brains with a graded series of MeOH:PBST washes at room temperature by incubating the specimens in 50:50 MeOH:PBST for 5 min, then in 30:70 MeOH:PBST for 5 min, and ending with 3 washes of PBST.
2. To permeabilize the brains, incubate them in PBST supplemented with proteinase K (PK) at a final concentration of 20 µg/ml PK for 30 min at 37 °C. Store aliquots of 10 mg/mL PK at -20 °C and thaw them before use.
3. Remove the PK solution and wash away any remaining enzyme by rinsing the brains with PBST 3 times over 15 min.
4. Refix the permeabilized brains by incubating them in 4% PFA for 20 min at 25 °C (or room temperature). Then, remove PFA and wash away any remaining fixative by rinsing the brains 3 times over 15 min with PBST.
5. Replace the final PBST rinse with freshly made immunoblocking (IB) buffer (10% normal goat serum and 1% DMSO in PBST) and incubate at room temperature for at least 1 h.
6. Dilute primary antibodies into IB buffer at the appropriate concentration (e.g., 1:500 for RFP and 1:300 for PH3 antibodies).
7. Remove IB buffer from brains and replace it with the primary antibody dilution. Incubate overnight at 4 °C on an orbital shaker with gentle rocking. Ensure tubes are securely resting on their sides.
8. The next morning, remove the primary antibody solution with a micropipette, rinse a couple of times in PBST, and then wash the brains in PBST at room temperature for 4 x 30 min.
   NOTE: Primary antibody dilutions can be reused once within ~7 days if stored at 4 °C.
9. Rinse the brains in IB buffer while diluting secondary antibodies into IB buffer at the appropriate concentration (e.g., 1:500 for Alexa-fluor 568 anti-rabbit).
10. Remove the IB buffer and replace it with the secondary antibody dilution. Incubate overnight at 4 °C on an orbital shaker with gentle rocking of the tubes resting on their sides.
11. The next morning, remove the secondary antibody solution, rinse with PBST, and then wash brains in PBST at room temperature for 4 x 30 min.
12. Counterstain with 4',6-diamidino-2-phenylindole (DAPI) or ToPro3 (1:5,000 in PBST at 4 °C overnight).
13. The next morning, transfer the larval brains into PBS and proceed to the mounting and imaging steps listed below.

10. Mounting and imaging

1. Secure a chambered slide into the lid of a 100 mm Petri dish with vacuum grease.
2. Transfer the immunostained and processed larvae to a well plate or depression slide to view them with a dissecting microscope.
3. Mount the larvae on the chambered slide in columns of 1% LMP agarose.
   1. Using a glass Pasteur pipette, put one larva on the chambered slide, depositing as little PBS as possible. With the same pipette, cover the larva with molten 1% LMP agarose (~40 °C).
   2. With a blunt tungsten needle or forceps, draw the agarose into a column and then position the larva as symmetrically as possible, with the dorsal surface visible.
   3. Repeat this procedure for the remaining larvae.
   4. Once the agarose has gelled, cover it with a few drops of PBS and then place it on the stage of the confocal microscope.
4. Align the objective lens with the sample, focus the microscope using transmitted light, and illuminate the sample with the appropriate lasers.
   NOTE: In this example, 405 nm and 568 nm wavelengths were used to excite DAPI and RFP, respectively.
5. Adjust the gain, offset, and laser power by moving the slider bars to an appropriate level relative to the signal and the used detector. Check the histograms for each channel and click the State of saturation indicator button above the image to gauge the level of exposure, ensuring that none of the pixels are saturated and that the signal-to-noise ratio is high. For an example of optimum confocal imaging parameters for cellular and subcellular resolution within zebrafish, see ²¹.
6. Set the upper and lower limits of the stack of z-planes by using the scrollwheel on the mouse to find the top and bottom of the sample. Click the top and bottom limits for image acquisition and then trigger the z-stack capture by clicking the Run now button.
   NOTE: Depending on how symmetrically the brain is mounted, the total z-depth for a 9 dpf larval zebrafish brain will be ~150-200 µm.
7. Process and colorize the images with colorblind accessibility in mind (e.g., use blue and orange or magenta and green for two-color images). Set lookup tables in the software used to capture the images or image processing software such as Photoshop from Adobe.
8. In Photoshop, color raw photomicrographs saved as 8 bit Tagged Image File Format (TIFF) by (i) converting the Image Mode to RGB and (ii) accessing the Curves option under the Image | Adjustments menus and then (iii) sliding the appropriate colored light outputs to 0 or 255 levels to achieve the desired color. For example, to achieve a green signal, select the Red channel and slide its output to 0; then, select the Blue channel and slide its output to 0 too. Similarly, to achieve a magenta signal, select the Green channel and slide its output to 0; leave the red and blue channels as they are.
   NOTE: Similar steps can be performed in the free Gnu Image Manipulation Program (GIMP).
9. Manually quantify the numbers of cells displaying a specific marker using the cell counter plugin in ImageJ²², available under Analyze in the Plugins menu. Examples of how to use ImageJ for cell counting can be found in many tutorials, including ²³.
10. Generate graphical representations of data in statistical software such as RStudio (see supplemental data for .Rmd file).

Representative Results

To confirm whether eye removal was complete and assess how the optic tectum changes, surgeries were performed in the Tg[atoh7:RFP] strain, which labels all RGCs with a membrane-targeted RFP and, thus, all axons that project from the retina and form the optic nerve²⁴. Although using this strain is not absolutely necessary, it enables straightforward observation and visualization of the optic nerve termini in the optic tectum neuropil. Other approaches for labeling the optic nerve, such as axon tracing with lipophilic DiI or DiO dyes^25,26 or multispectral labeling with brainbow²⁷, would also be possible.

As shown in Figure 2, progressive degeneration of retinal axons in the optic tectum neuropil was observed following eye removal. By two days post surgery, RFP-labeled axons exhibited hallmarks of rapid Wallerian degeneration²⁸ such as blebbing and fragmentation (Figure 2A,B). Recent work has shown that microglia engulf more myelin material when neurons are silenced²⁹. Consistent with microglia engulfing bits of the degenerating RGC axons and migrating away from the source of degeneration, bright RFP-labeled puncta within and outside the tectal neuropil were noted (Figure 1B, arrows). By four days post surgery, fragmented axons and RFP-labeled axonal debris are considerably reduced in either tectal lobe, indicating relatively rapid clearance of the dying and degenerating axons (Figure 2C,D).

To perform wholemount immunohistochemistry and visualize the optic tectum of larval zebrafish, the brain was exposed by dissecting off the eye(s), jaw, ears, and skull cap of the skin and connective tissues. Ideally, the brain remains intact during this procedure (e.g., Figure 2C,D). However, parts of the forebrain, particularly the olfactory bulb, are likely to be damaged or completely removed when the skull cap of skin or jaw is removed (Figure 2A, arrowhead). Moreover, sometimes the lateral edge of the tectum can be sliced when piercing or pinching the skin to pull it off the brain (e.g., Figure 2A; tear on the right tectal lobe).

Immunohistochemistry was also used to assess the number of mitotic cells in innervated and non-innervated lobes of the optic tectum by staining whole brains with a PH3 antibody. These data show significantly fewer mitotic cells on the denervated side of one-eyed larval fish (p = 0.00033, Welch's t-test; Figure 3).

Figure 2: Eye removal at 5 dpf triggers degeneration of optic nerve axons. (A-D) Representative maximum-intensity projections showing dorsal views of wild-type brains from intact larvae fixed at 7 dpf (A) or 9 dpf (C), or one-eyed larvae from eye extirpation surgeries at 5 dpf, fixed 2 dps (B) or 4 dps (D). All nuclei are stained with DAPI (green), and optic nerve axons terminating in the neuropil of the optic tectum are labeled by the atoh7:RFP transgene (magenta). Asterisk indicates innervated tectal neuropil in larvae with only the right eye intact. Arrows indicate examples of fragments of degenerated RGC axons emanating from the denervated optic tectum. Arrowhead indicates missing parts of the forebrain. Scale bar = 50 µm. Abbreviations: dpf = days post fertilization; dps = days post surgery; RGC = retinal ganglion cells; DAPI = 4',6-diamidino-2-phenylindole. Please click here to view a larger version of this figure.

Figure 3: Number of mitotic cells in the optic tectum increases with innervation. (A-B) Representative maximum-intensity projections showing dorsal views of wild-type brains from intact (A) or one-eyed (B) 9 dpf larvae immunostained with antibodies for the mitotic marker PH3 (magenta) and nuclei counterstained with DAPI (green). Asterisk indicates innervated tectal neuropil from the intact right eye. (C) Box plot with all individual data points overlaid showing quantitation of the ratio of PH3⁺ cells in innervated (left) versus denervated (right) tectal lobe as a difference ratio (left-right/total) for one-eyed (n = 12) and two-eyed (n = 8) 9 dpf larvae. Means of the two conditions compared with Welch's t-test (***, p < 0.0005). Scale bar = 50 µm. Abbreviations: dpf = days post fertilization; dps = days post surgery; PH3 = phosphorylated histone H3; DAPI = 4',6-diamidino-2-phenylindole. Please click here to view a larger version of this figure.

Supplemental file: Outline and instructions for plot shown in Figure 3C. This file contains all data and code so that anyone can reproduce the box plot with all data points overlaid, as shown in Figure 3C. Please click here to download this File.

Discussion

The techniques described in this paper illustrate one of many approaches for studying vertebrate visual system development in zebrafish. Other researchers have published methods to dissect the embryonic retina and perform gene expression analyses¹⁹ or visualize neuronal activity in the optic tectum³⁰. This paper provides an approach for exploring how differential retinal input may influence cell behaviors in the optic tectum.

To ensure successful eye extirpation and larval survival after surgery, it is important that the larvae are healthy, adequately anesthetized, and immobilized in 1% agarose. It is also crucial that forceps and tungsten needles are very sharp and clean. Larvae survive best when they are not fed 24 h before or after the surgery, and rearing larvae in MMR supplemented with antibiotics speeds the healing process. Importantly, the higher ionic strength of MMR and especially the higher concentration of calcium promotes healing¹⁴. However, longer exposure to the higher salinity can decrease survival, similar to what has been documented in embryonic zebrafish³¹. A most critical step is fixation to ensure successful brain dissections, which are essential for data analysis of images of immunostained samples. Using freshly made 4% PFA supplemented with 4% sucrose (affectionately called sweet fix) tends to increase the ease of skin removal and brain tissue preservation. As with the surgeries, sharp and clean tools are a must for dissecting brains.

One of the biggest challenges of this protocol is embedding larvae for eye extirpations. It is important that each person find the way that works best for them so that they can confidently and quickly remove the larval eye. If the larva is accidentally stabbed during the surgery, humanely kill it by decapitation. Another challenge is determining the most effective antibody concentrations for wholemount immunostaining. When determining the appropriate antibody concentrations, use the published literature as a starting point and optimize the protocol as using published reagents on different tissues (especially on larger tissues or older animals) may require increased concentrations and/or incubation times.

The major limitation of this method is the time a single experiment takes from start to finish and the number of fish that can be studied at any given time. For example, all the data shown in Figure 3 took about one month from the initial pairing of fish to the final data collection and analysis. This relatively low-throughput approach could be complemented by heterologous genetic approaches that either selectively silence neural activity in the visual system^32,33 or by mutants that lack RGC axons or activity^34,35.

The significance of this method is that it brings a traditional embryological technique to bear on modern transgenic fish lines, allowing direct comparison of cellular and genetic mechanisms operating in each of the tectal lobes within the same individual. Moreover, this method is ripe for application to other experimental systems, including Astyanax surface fish³⁶ and/or transgenic zebrafish larvae with fluorescently-labeled microglia^37,38,39 and astrocytes⁴⁰ to study how the developing fish nervous system responds to such a dramatic injury.

Materials

Name	Company	Catalog Number	Comments
Equipment and supplies:
Breeding boxes	Aquaneering	ZHCT100
Dow Corning high vacuum grease	Sigma or equivalent supplier	Z273554
Erlenmeyer flasks (125 mL)			For making Marc's Modified Ringers (MMR) with antibiotics for post-surgery incubation.
Fine forceps - Dumont #5	Fine Science Tools (FST)	11252-20
Glass Pasteur pipettes	DWK Lifescience	63A53 & 63A53WT	For pipetting embryos and larvae.
Glass slides for microscopy	VWR or equivalent supplier	48311-703	Standard glass microscope slides can be ordered from many different laboratory suppliers.
Glassware including graduated bottles and graduated cylinders			For making and storing solutions.
2-part epoxy resin	ACE Hardware or other equivalent supplier of Gorilla Glue or equivalent	0.85 oz syringe	https://www.acehardware.com/departments/paint-and-supplies/tape-glues-and-adhesives/glues-and-epoxy/1590793
Microcentrifuge tube (1.7 mL)	VWR or equivalent supplier	22234-046
Nickel plated pin holder (17 cm length)	Fine Science Tools (FST)	26018-17	To hold tungsten wire while sharpening and performing surgeries/dissections.
Nylon mesh tea strainer or equivalent	Ali Express or equivalent		For harvesting zebrafish eggs after spawning; https://www.aliexpress.com/item/1005002219569756.html
Paper clip			For Tungsten needle sharpening device.
Petri dishes 100 mm	Fischer Scientific or equivalent supplier	50-190-0267
Petri dishes 35 mm	Fischer Scientific or equivalent supplier	08-757-100A
Pipette pump	SP Bel-Art or equivalent	F37898-0000
Potassium hydroxide (KOH)	Sigma	909122	For Tungsten needle sharpening device. Make a 10% w/v solution of KOH in the hood by adding pellets to deionized water.
Power supply (variable voltage)			For Tungsten needle sharpening device. Any power supply with variable voltage will work (even one used for gel electrophoresis).
Sylgard 184 Elastomer kit	Dow Corning	3097358
Tungsten wire (0.125 mm diameter)	World Precision Instruments (WPI)	TGW0515	Sharpen to remove eye and dissect larvae.
Variable temperature heat block	The Lab Depot or equivalent supplier	BSH1001 or BSH1002	Set to 40-42 °C ahead of experiments.
Wide-mouth glass jar with lid (e.g., clean jam or salsa jar)			For Tungsten needle sharpening device.
Wires with alligator clip leads			For Tungsten needle sharpening device.
Microscopes:
Dissecting microscope			Any type will work but having adjustable transmitted light on a mirrored base is preferred.
Laser scanning confocal microscope			High NA, 20-25x water dipping objective lens is recommended. Microscope control and image capture software (NIS-Elements by Nikon) is used here but any confocal microscope will work.
Reagents for surgeries and dissections:
Calcium chloride dihydrate	Sigma	C7902	For Marc's Modified Ringers (MMR) and embryo medium (E3).
HEPES	Sigma	H7006	For Marc's Modified Ringers (MMR).
Low melting point agarose	Invitrogen	16520-050	Make 1% in embryo medium (E3) or Marc's Modified Ringers (MMR).
Magnesium chloride hexahydrate	Sigma	1374248	For embryo medium (E3).
Magnesium sulfate	Sigma	M7506	For Marc's Modified Ringers (MMR).
Paraformaldehyde	Electron Microscopy Sciences	19210	Dilute 8% (w/v) stock with 2x concentrated PBS (diluted from 10x PBS stock).
Penicillin/Streptomycin	Sigma	P4333-20ML	Dilute 1:100 in Marc's Modified Ringers.
Phosphate buffered saline (PBS) tablets	Diagnostic BioSystems	DMR E404-01	Make 10x stock in deionized water, autoclave and store at room temperature. Dilute to 1x working concentration.
Potassium chloride	Sigma	P3911	For Marc's Modified Ringers (MMR) and embryo medium (E3).
Sodium chloride	Sigma	S9888	For Marc's Modified Ringers (MMR) and embryo medium (E3).
Sodium hydroxide	Sigma	S5881	Make 10 M and use to adjust pH of MMR to 7.4.
Sucrose	Sigma	S9378
Tricaine-S	Pentair	100G #TRS1	Recipe: https://zfin.atlassian.net/wiki/spaces/prot/pages/362220023/TRICAINE
Reagents for immunohistochemistry:
Alexafluor 568 tagged Secondary antibody to detect rabbit IgG	Invitrogen	A-11011	Use at 1:500 dilution for wholemount immunohistochemistry.
DAPI or ToPro3	Invitrogen	1306 or T3605	Make up 1 mg/mL solutions in DMSO; 1:5,000 dilution for counterstaining.
Dimethyl sulfoxide (DMSO)	Sigma	D8418	A component of immunoblock buffer.
Methanol (MeOH)	Sigma	34860	Mixing MeOH with aqueous solutions like PBST is exothermic. Make the MeOH/PBST solutions at least several hours ahead of time or cool them on ice before using.
Normal goat serum	ThermoFisher Scientific	50-062Z	A component of immunoblock buffer. Can be aliquoted in 1-10 mL volumes and stored at -20 °C.
Primary antibody to detect phosphohistone H3	Millipore	06-570	Use at 1:300 dilution for wholemount immunohistochemistry.
Primary antibody to detect Red Fluorescent Protein (RFP; detects dsRed derivatives)	MBL International	PM005	Use at 1:500 dilution for wholemount immunohistochemistry.
Proteinase K (PK)	Sigma	P2308-10MG	Make up 10 mg/mL stock solutions in PBS and use at 10 µg/mL.
Triton X-100	Sigma	T8787	Useful to make a 20% (v/v) stock solution in PBS.
Software for data analysis
ImageJ (Fiji)			Freeware for image analysis; https://imagej.net/software/fiji/
RStudio			Freeware for statistical analysis and data visualization; https://www.rstudio.com/products/rstudio/download/
Adobe Photoshop or GIMP			Proprietary image processing software (Adobe Photoshop and Illustrator) are often used to compose figures). A freeware alternative is Gnu Image Manipulation Program (GIMP; https://www.gimp.org/)
Zebrafish strains. This study used the AB, TU, Tg[atoh7:RFP] strains.			Available from the  Zebrafish International Resource Centers in the US (https://zebrafish.org/home/guide.php) or in Europe (https://www.ezrc.kit.edu/). Specialized transgenic strains that have not yet been deposited in either resource center can be requested from individual labs after publication.