Astyanax breeding has been described elsewhere (Elipot et al. 2014; Peuß et al. 2019; Ma et al. 2021). Briefly, an increase of temperature from 22 °C to 26.5 °C in the first three days of a breeding week provides two-to-three consecutive spawning days with maximal numbers of high-quality embryos, which is then followed by a decrease of temperature from 26.5 °C to 22 °C during the last three days of the spawning week. Most spawning occurs at night for both surface and cavefish. In the morning, collect embryos in 100 × 15 mm Petri dishes, sort and clean them, and transfer them into embryo medium (EM-Westerfield 2000) with methylene blue. An alternative is to conduct in-vitro fertilization.

Keep the embryos in EM in a 23 °C incubator until the desired stage. The lens becomes visible in Astyanax kept at 23 °C at 18.5 hpf. Hatching occurs at 24.5–28 hpf. The lens enters apoptosis at about 25 hpf. When conducting lensectomy on surface fish to replicate the effects of lens degeneration in cavefish, the optimum time is 1–3 dpf, or within 48 hrs after hatching. If lensectomy is to be conducted before hatching, remove the chorion manually with two pairs of sharp forceps, and incubate the embryos in 0.2% EDTA in Calcium-free Zebrafish Ringer’s (ZFR) for 30 minutes.

In previous protocols, two needles were used. One with a blunt tip needle made of a thin tungsten wire and a second one with a sharp tip made by holding the tungsten wire over a Bunsen burner for 1–1.5 minutes, burning off the metal, and creating a very fine tip). Previous protocols instructed lensectomies to be conducted by hand under a microscope. Since the lens is only about 50 μm, extreme precision is required to ablate the lens without harming other structures. Normal tremor of the hands makes this extremely challenging, even for highly trained people.

In this improved protocol, instead of using tungsten needles held by hand, microinjection needles were made from glass capillaries mainly with a Narishige’s PC-10 Dual-Stage Glass Micropipette Puller, with the puller was set to a one step weighted pull at 70.5 °C. Other brand micropipette pullers were tested and found to give similar results. Borosilicate glass capillaries are heated and pulled to get extremely fine and sharp needles, similar to those used for cell injections (Fig. 1A, B). These needles are then attached to a manual micromanipulator (Type MM33 Rechts; Märzhäuser, Wetzlar, Germany). The micromanipulator allows for precise movements in the X, Y, and Z directions, eliminating hand tremor effects and maximizing the precision of movements. We found that the best technique was with one hand controlling the micromanipulator, while the other hand moved the petri dish that holds the specimen (Fig. 1C). This combination allowed for the best combination of dexterity and efficiency of movements.

Figure 1. 

A for the preparation of dissection needles, microinjection needles are made from borosilicate glass capillaries with a Micropipette Puller B glass capillaries are heated and pulled to get extremely fine and sharp needles C instead of manipulating the dissection needle by hand, a micromanipulator is used. This dramatically reduces jittery movements that can puncture neighboring structures such as the brain or the heart. The micromanipulator allows precise puncturing around the lens with the needle’s movements controlled easily at less than 5 μm D clean needles are essential. Throughout the procedure, the needle progressively gets covered with a fatty substance that essentially blunts the needle. It is best to exchange it for a new one E embryo a week after a one-sided lensectomy in dorsal view. The left, lensectomized eye is regressing.

• Embryo medium (EM; pH 7.0, per liter, contains: 10 ml Hanks Solution #1, 1 ml Hanks Solution #2, 10ml Hanks Solution #4, 10 ml Hanks Solution #5, 0.35 g sodium bicarbonate, 300 uL of 2M HCl, penicillin-streptomycin 500,000U) As in The Zebrafish Book (University of Oregon Press, 2000).
• Anesthetizing solution:10 ml EM, 1 mg ms-222 (tricaine methanesulfonate), 1 mg NaHCO _3.
• 2% low melting agarose: 10 ml EM, 1 mg ms-222 (tricaine methanesulfonate), 1 mg NaHCO ₃, 0.2 gr low melting agarose.
• Pulled glass capillaries.
• Dissection needle with a thin tungsten wire blunt tip.
• Mini scalpel.
• Plastic pipette dropper (wide mouth so the embryo can easily fit).
• 200 µl pipette.
• Petri dishes.
• Dissection microscope.
• Manual micromanipulator.

• At the desired stage (40 hpf, for example), take the embryos kept in EM in the incubator. With a plastic pipette dropper, carefully transfer the embryos to the anesthetizing solution and incubate for about 30 seconds or until the embryos stop moving. The number of embryos depends on proficiency. We operated up to ten specimens simultaneously, but for best survival and efficiency, about three at a time is recommended.
• With the pipette dropper, transfer anesthetized specimens to a petri dish (Fig. 2A). Before adding the agarose, absorb the excess liquid (Fig. 2B). It was found that, otherwise, the agarose surrounding the embryo would be diluted and would not grasp as firmly during the micromanipulations. However, caution should be used that the embryo does not dry before adding the agarose.
• Add the 2% low melting agarose when the melted agarose is close to room temperature but before it solidifies on and around the embryo (Fig. 2C). Most embryos will naturally lie on their sides in the agarose. Reorient those that do not lie in this position so that one eye is facing up before the agarose solidifies. The depth at which the embryo lies within the agarose drastically changes the performance. When very deep, it is difficult to see the structures, and when traversing the agar, the needle gets deflected from its target. When barely covered, it risks detaching the specimen from the agar. We found that adding about 100 μl agarose worked well, but sometimes excess agarose must be reabsorbed when the specimen sinks or is attached to the petri dish.
• Wait for the 2% agarose to solidify. For time efficiency, the Petri dish containing the samples can be put in a refrigerator for 1–2 min. It was also found convenient to use this time to make a pair of microinjection needles. After the agar solidifies, the Petri dish is transferred under a dissecting microscope (Fig. 1C). Microscopes of high quality, good resolution, and good illumination are recommended for the best results. By its side, there will be the manual micromanipulator with the microinjection needle attached to it (Fig. 1C). Adding a few drops of liquid anesthetizing solution on top of the agarose is optional. Depending on the specific sample, it can enhance or reduce visual clarity.
• Using the microinjection needle, very carefully cut the lens out from the embryos using small strokes. Make sure to cut just close enough to the lens that it neither tears it, nor removes too much tissue from the rest of the eye. We developed three styles, each with its advantages and disadvantages:

Style 1: With the micromanipulator, slowly bring down the needle by the side of the lens. Pressure down until it makes a puncture in the surface ectoderm/cornea at the junction between the lens and optic cup (Fig. 3A). Depending on the specimen and how solid the agarose grabs the specimen, the eye can deform and sink slightly down with the pressure until the needle ruptures the tissue. Be careful that the needle does not go further down and punctures other structures, such as the brain or the heart. The use of the micromanipulator is a great improvement to the previous protocols that used a hand-held needle in this regard. After a hole is made with the micromanipulator, pull the needle out. With the hand holding the petri dish, slightly reposition the sample so the needle can make a different puncture at another site near the lens (Fig. 3B). Repeat these punctures with a circling pattern around the lens. Ten to fifteen punctures are performed (Fig. 3C). Afterward, with a coordinated motion of the hand holding the petri dish and the micromanipulator, insert the needle between punctures and gently pull the needle out to tear the tissue between punctures. Repeat this progressive tearing of tissue around the lens. Finally, put the needle on one side, under the lens, and push it out of the optic cup (Fig. 3D). It is common for the lens to brake during this step, especially if no liquid has been added above the agar. With the needle, any lens remains can then be scooped out.

Figure 2. 

A specimens are transferred to a Petri dish after being in anesthetizing solution for 30 seconds or until embryos stop moving B absorb the excess liquid C add 2% agarose EM. The depth at which the embryo lies should not be too deep because the needle gets deflected from the target, and it is difficult to see the structures. However, it risks detaching the specimen from the agar when barely covered. After agarose solidifies, proceed with lensectomy D–G if lensectomy is to be done on both sides, with a scalpel, cut a rectangle of the agar around the specimen. Very gently slide the scalpel under the rectangle of agar with the specimen. Helped with twicers, flip around the agar slab, so the specimen is on the other side. More than one specimen can be done at a time to increase yield H add 2% agarose around the rectangle of agar or over the specimen if it was dislodged I after the second lensectomy is done, submerge in embryo media and gently dislodge the embryo from the agar with downward strokes starting around the tail and ending on the head.

Figure 3. 

Style #1 for doing lens ablations A with the micromanipulator, slowly bring down the needle by the side of the lens and pressure down until it makes a perforation on the surface ectoderm/cornea B repeat these punctures around the lens C insert the needle between punctures, and with a coordinated motion of the hand holding the petri dish and the micromanipulator, gently pull the needle out to tear the tissue between punctures D put the needle on one side, under the lens and push it out of the optic cup.

Style 2: With the micromanipulator, slowly bring down the needle by the side of the lens and puncture the surface ectoderm. While inserted, position the needle so that it is under the lens (Fig. 4A). Afterward, move the needle away from the eye with the lens positioned in the center. This is done with a coordinated motion of the hand holding the Petri dish and the micromanipulator. The optic cup will distend until it rips open, with the lens bursting out of the eye cup (Fig. 4B). In most cases, the lens will be obliterated. With the needle, any lens remains can then be scooped out. Style #2 is riskier because, depending on the robustness of the specimen and the grip the agarose has on the specimen, if the ripping of the tissue is done too fast, it can dislodge other internal tissues and kill the specimen. If the agar cover is not too deep, the specimen can also detach from the agar before the lens is ripped out. We found that experience and gentle slowness while pulling was necessary to perform this style effectively. Once this style is mastered, time dedicated per lensectomy is shortened without significantly reducing survival.

Figure 4. 

Style #2 for doing lens ablations A with the micromanipulator, slowly bring down the needle by the side of the lens and puncture the surface ectoderm. While inside the optic cup, position the needle so that it is under the lens B with a coordinated motion of the hand holding the Petri dish and the micromanipulator, move the needle away from the eye with the lens position in the center. The optic cup will distend until it rips open, with the lens bursting out of the eye cup.

Style 3: With the micromanipulator, slowly bring down the needle just by the side of the lens, closer than the previous two styles, and puncture the overlying ectoderm. Position the needle so that it is above the lens instead of below (Fig. 5A). Gently scrape the overlaying agarose and scrape the tissue over the lens. The tissue will tear, and the lens will float up if there is liquid (Fig. 5B). It may need some nudging with the needle.

Figure 5. 

Style #3 for doing lens ablations A with the micromanipulator, slowly bring down the needle just by the side of the lens, closer than the previous two styles, and puncture the surface ectoderm. While inside the optic cup, position the needle so that it is above the lens instead of below. Gently scrape the overlaying agarose and scrape the tissue over the lens B the tissue will tear, and the lens will float up if there is liquid. It may need some nudging with the needle.

• If lensectomy is to be done on only one side of the face, submerge the agar containing the specimen in embryo medium and gently dislodge the embryo from the agar; with a blunt tip dissection needle made with a thin tungsten wire or the hair of a toothbrush, stroke downward the agar around the embryo starting around the tail. Pieces of agar will dislodge. Progressively dislodge fragments of agar from tail to head until the specimen is released (Fig. 2I). Caution should be used to minimize touching the specimen, especially with the tip of the needle. Alternatively, fine-tip twicers can dislodge agar on either side of the embryo, again, from tail to head. Once free, the embryo should be transferred to clean embryo medium and put in the incubator.
• If lensectomy is done on both sides, it is best to have the agar dry, not overlaid with liquid. With a scalpel, cut a rectangle of the agar around the specimen (Fig. 2D). Very gently slide the scalpel under the rectangle of agar with the specimen. Helped with twicers, flipping around the agar slab, to expose the specimen on the other side (Fig. 2E, F). If the specimen is dislodged from the agar, add embryo medium to keep it under water. Gently flip it around. Flipping around specimens that have dislodged from the agar slab involves directly contacting the specimen and tends to cause more harm.
• Add 2% agarose around the rectangle of agar or over the specimen if it was dislodged (Fig. 2G, H). Caution should be used in not adding too much agar above the head of the specimen. When too deep, the needle gets deflected from its target, and it is difficult to see the structures.
• Repeat the previous steps to dislodge the second lens.

Style #3 works best in younger embryos that have just hatched. In older specimens, the tissue covering the lens has grown, which may require the stronger tearing of styles #1 or #2. Style #3 is the preferred style of lens removal if lenses are to be collected for transplants or other studies.

In this case, at the beginning of the protocol, after embedding the specimen in 2% agarose, overlay it with EM containing 1.2% agarose. Use style #3 preferentially. Once free, lenses will float in the medium. Using the blunt needle, carefully push the lens of the donor to just above where the host lens would normally be, and then push it down into the eye with the blunt needle. The host lens may be discarded.

Leave donor and host embryos in 1.2% agarose for 30–60 minutes, then release them from the agarose using the sharp needle and transfer them into EM in the incubator.

Healthy conditions for the breeding colony: Survivorship of embryos can be drastically different between laboratories due to the conditions in which parents and the embryos are kept. The technique described here can produce live embryos, with the limiting factor being the general survivorship of embryos within the specific laboratory conditions they are kept.

Clean needles: Throughout the procedure, the needles progressively get covered in what appears to be a fatty substance (Fig. 1D). This “glob” blunts the needle, and instead of easily puncturing the tissue, it pushes down the eye until uncontrolled rupture may happen. Sometimes the needle tip can be cleaned by immersion in the agar and sliding the needle sideways while inside the agar. Nonetheless, it is best to exchange it for a new one after about three lenses have been removed, or when the glob develops.

The agar’s depth significantly affects the efficiency of the procedure (Fig. 2C). Especially when doing lens ablation on both sides, one side will be overlaid by more agar. Practice until achieving correct conditions. We found that, as a generality, adding too much agar was a more significant problem.

Gentle, slow motions are to be done with the micromanipulator throughout the process (Fig. 1C). While up to 10 fish were successfully put in a row to undergo the procedure, we found that a slower approach, working with about three fish at a time, was the most efficient (Fig. 2G–I).