A 3-D Tail Explant Culture to Study Vertebrate Segmentation in Zebrafish

This protocol involves use of live vertebrate embryos younger than 1 day post-fertilization. All the animal experiments were performed under the ethical guidelines of Cincinnati Children's Hospital Medical Center; animal protocols were reviewed and approved by the Institutional Animal Care and Use Committee (Protocol # 2017-0048).

1. Embryo collection

1. Set up pairs of zebrafish in crossing tanks the night before the embryo collection day. For precise staging control of embryo development, use barriers between mating pairs.
2. Raise barriers before preferred spawning time and collect eggs within 15 minutes in a 100 mm Petri dish.
   1. Clean debris from the Petri dish. If more than 50 embryos are collected from a single clutch, split the clutch into multiple Petri dishes accordingly.
3. Incubate embryos in fish system water at 28 °C until they reach 50% epiboly stage (5 hours post-fertilization). A standardized embryo growth medium such as E3 can also be used instead of the aquarium system water until step 3.2.
   1. Remove unfertilized eggs under a stereoscope and move the embryos to a 23.5 °C incubator overnight (O/N). Embryos should be at 8-10 somites stage the morning following collection day.

2. Tool preparation

1. Sterilize the microsurgery knife blade, needle tips (used for dissection of the tissue), and glass Pasteur pipette by soaking in 100% ethanol (EtOH) and fire glazing.
2. Use two layers of transparent tape (~100-120 µm thickness) on 25 mm x 75 mm microscope slides. Cut ~18 mm x 18 mm square wells in the center of each slide's tape covering with a scalpel.
   1. Wipe the prepared slide chambers with 70% EtOH. These wells will hold ~40 µL of medium.

3. Sample preparation

1. Dechorionate embryos using the tip of two needle syringes under a stereoscope. Transfer embryos into a separate Petri dish with fish system water to rinse.
2. Using a fire-glazed sterile glass Pasteur pipette, transfer embryos in a 6 cm Petri dish containing dissection medium (Leibovitz-15 cell culture medium with L-Glutamine without Phenol Red, 0.8 mM CaCl₂ and 1× antibiotic-antimycotic solution).
   NOTE: Continue to use a sterilized glass pipette for all transfers following this step.
   1. Use a glass Petri dish for explanting procedure to avoid polystyrene chips during dissection.
3. Put 50 µL of tissue growth medium (dissection medium, and 10% FBS) in the slide chamber.
4. Stabilize an embryo for dissection under the stereoscope with a needle tip at the yolk-tissue intersection near the hindbrain.
5. Keeping the embryonic tissue stable with a needle, use the microsurgical knife with the blade held at 45° to cut the tissue anterior to the hindbrain apart and peel the yolk off the embryonic tissue starting from the anterior and moving towards the tail bud (Figure 1A).
   NOTE: Be careful to not lose the skin tissue while cleaning the yolk. The skin would easily peel off as a flanking single layer elastic tissue around the embryo during the dissection, so it is easy to recognize.
6. Once the yolk is fully removed from the embryonic body, cut the flanking skin tissue from the tailbud. Keeping the last formed 3-4 somites intact, cut the more anterior tissue out (full-axis explant).
   1. The yolk should come off mainly intact from this procedure. In case of a ruptured yolk, significant granules of yolk can stay attached to the ventral surface of the tissue. If so, use a lash tool to clean off remaining yolk granules gently.
      NOTE: An imbalance of skin tissues on the lateral sides of an explant would not allow the tissue to maintain a straight growth direction. The explant instead will bend towards the side of more stretched skin. This imbalance can be corrected under the stereoscope by rupturing the skin layer with the aid of microsurgical knife.
   2. For skinless explants, press on a tip of the skin layer with needle and peel the tissue explant off with the microsurgical knife. These explants will not elongate their body axis in culture.
   3. In addition to the full axis explants, alternative explants can be made at this step. For instance, dissect out already segmented somites using the microsurgical knife (full-PSM explants) or dissect the PSM into its half anteroposterior (half-PSM explants). Please see section 5.1 for an application of such alternative explants.
7. Immediately transfer the dissected explant to a 22 mm x 22 mm coverslip on which the imaging will be performed.
   1. Arrange the explant flat on the dorsoventral axis, ventral side touching the coverslip (Figure 1B). Gently remove the excess media around the tissue explant using a 20 µL filtered tip pipette.
      NOTE: Delayed transfer of dissected explants to the coverslip results in deformations of the tissue, as it is relieved from the geometric constraints of the yolk.
8. Swiftly and carefully flip the coverslip with the explant over the growth medium filled slide chamber.
   1. To prevent bubble formation, place a side of the square coverslip on the tape chamber and release the other side gently. Take care to not move/deform the explant in this step.
   2. Gently remove excess media bleeding out of the chamber by pressing the slide chamber on a laboratory tissue. The coverslip will sit stably on the slide chamber for live imaging, due to the surface tension of the liquid medium without any sealing.
   3. For long term culturing (>6 hours), use a bigger chamber. 22 mm x 50 mm rectangular coverslips together with two parallel lanes of tape layers on slides can be utilized in such cases. A ~1 mm wide gap can be left in between two tape lanes to facilitate air access into the growth medium.
9. Repeat steps 3.3-3.8 to prepare more explants. Prepared explants will elongate their A-P body axis with an average of ~30 µm/h speed and segment their somites with ~40 min intervals at 25 °C (Figure 2A, Video 1).
   1. For non-elongating explants, apply gentle pressure to the sides of slide holding the sample in step 3.8.2 while sucking the excess media out on a laboratory tissue. Alternatively, explants can be cultured in single tape layer slide chambers. Also, chemically activating the slide chamber surface with Type I Collagen will result in non-elongating explants (Figure 2B, Video 2).
      1. Perform coating of the chamber with Type I Collagen in advance by fully covering the slide chambers with 15-20 mL of prediluted collagen solution at room temperature for 1 h. Use a laminar flow hood for this protocol to maintain sterility. Carefully rinse chambers with dissection medium at the end.
   2. For embryos older than 15 somite stage, mount the tail explant tissue laterally instead of a flat (dorsoventral) mount (Video 3). To prevent muscular twitching, include 0.004% tricaine solution in the culture media as an anesthetic agent³.

4. Live image acquisition

1. Image samples either on a dissection scope for widefield transmitted light imaging of somite segmentation sizes and periods, or with structured illumination/confocal/light sheet microscopy using transgenic reporter fish lines.
2. Equilibrate the temperature of tissue explants with the imaging room temperature for at least 15 min.
   1. For more precise temperature control, use a commercial temperature control system mounted on an inverted microscope.
3. Set the image acquisition frame intervals to 2 – 10 min depending on the biological process of interest.
   NOTE: Zebrafish somite segmentation is a fast process, ranging between 20 – 55 min for viable temperatures of 30 °C to 21.5 °C in whole embryos. Explants will elongate and segment ~30% slower than the whole embryos.
   1. Pay attention to leave enough delay between sets of channel acquisitions to avoid possible phototoxicity to live tissue. Do not expose the tissue to the excitation beam for more than half of the imaging duration and lower the beam intensity as much as possible.
      NOTE: Accumulation of reactive oxygen species (ROS) is generally the major cause of phototoxicity in live samples⁴. Ascorbic acid as a ROS scavenger can be supplemented to growth medium at 4 mM concentration to buffer ROS activity and alleviate phototoxicity. Adverse effects of phototoxicity might be hard to notice during live imaging. Tail explants are advantageous in this aspect since some visual markers of phototoxicity such as mitotic arrest, impeded tissue development (i.e., formation of somites, tail elongation), and disintegrating tissue are easier to notice. Please refer to the provided reference⁴ for a detailed discussion.
4. Use single cell-stage RNA injected embryos to acquire 4-D images intended to be segmented and analyzed in cellular resolution level.
   1. Use 300 pg of RNA from in vitro transcribed membranes and nuclear fluorescent reporter marker plasmids such as pCS-membrane-ceruleanFP (Addgene plasmid #53749) or pCS-memb-mCherry (Addgene plasmid #53750) in combination with pCS2+ H2B-mTagBFP2 (Addgene plasmid #99267) or pCS2+ H2B-TagRFP-T (Addgene plasmid #99271) in injections. For a sample movie with cell membrane and nuclei markers please see Video 4.
      ​NOTE: Average cell size of PSM tissue is about ~5 µm in diameter, of which nuclei comprises ~3 – 4 µm. A pixel size of ~0.5 µm and a z-sectioning of ~1 µm should be recorded for proper cell segmentation.

5. Immunostaining of tail explants

NOTE: Tissues grown after various dissection scenarios (elongating, non-elongating, tail bud dissected, half PSM etc.) as flat-mounted tail explants⁵ can be recovered from slide chambers for further immunostaining quantifications of proteins of interest. Here, we present the protocol used for di-phosphorylated extracellular signal regulated kinase (ppERK) staining of explants as FGF signaling gradient readout.

1. After formation of somites until the desired stage, cautiously shift the coverslip halfway to the corner of the slide chamber without lifting.
2. With the help of ~100 µL of supplementary dissection medium in a glass Pasteur pipette, recover the explants from the slide and transfer into a 64-well cell culture plate.
   NOTE: Beginning with this step, all solution replacements can be performed under a dissection scope with a separate glass pipette for fixed samples. This will ensure not losing explant tissues in wells or transferring them in between.
3. After transferring all explants, suck the excessive medium out of the wells one-by-one and put 100 µL of 4% paraformaldehyde in PBS (PFA) into each well.
   CAUTION: PFA is a toxic solution with carcinogenic effects. Proper PPE should be used while handling.
4. Fix explants in a 64-well plate at room temperature for 1 h on a shaker.
   1. Tissue explants are more sensitive to deformations than whole embryos. Adjust the shaker speed accordingly.
5. Wash the fixative out with 150 µL of PBS-Tw (0.1% Tween20 in PBS) three times. Collect the first wash in a specific "PFA Waste" container.
6. Dehydrate explants in 4×5 min steps by replacing ~40 µL of solution each time with 100% methanol (MeOH).
   CAUTION: MeOH is a toxic chemical which is volatile and flammable. Work in a well-ventilated space and use proper PPE for handling.
7. As the last step of dehydration, remove all solution from wells and replace with 100 µL of MeOH. Incubate at -20 °C for 15 min.
   NOTE: Use a specific "MeOH Waste" container to collect the solutions until step 5.11.
8. Add 50 µL of MeOH and shake at room temperature for 5 min.
9. Rehydrate explants in 4×5 min steps by replacing ~40 µL of solution each time with PBS-T (0.1% Triton-X 100 in PBS). Use a specific "MeOH Waste" container to collect the solutions.
10. As the last step of rehydration, remove all solution from wells and replace with 100 µL of PBS-T.
11. For tissue permeabilization treat explants with 1.5% Triton-X 100 in PBS for 20 min at room temperature on the shaker.
12. Wash samples with MAB-D-T (0.1% Triton-X 100 detergent and 1% dimethyl sulfoxide (DMSO) in 150 mM NaCl 100 mM maleic acid buffer pH 7.5) 3×5 min.
    CAUTION: DMSO is flammable and a toxic mutagen. Proper PPE should be used while handling.
13. Incubate explants in 100 µL/well serum blocking solution (2% fetal bovine serum in MAB-D-T) for 2 hours at room temperature.
14. Replace all the blocking solution with 50-100 µL/well primary antibody solution (1:1000 dilution of monoclonal mouse antibody against ppERK in serum block). Incubate samples O/N (>16 h) at 4 °C on shaker.
15. Wash the primary antibody solution with MAB-D-T 5×5 min.
16. Incubate samples in secondary antibody solution (Alexa Fluor 597 goat anti-mouse IgG2b (1:200) and Hoechst 33342 (1:5000) in MAB-D-T) O/N at 4 °C on a shaker or for 3 h at room temperature.
    NOTE: Beginning at this step, cover the 64-well plate with aluminum foil to avoid light exposure of secondary antibody treated samples.
    CAUTION: Hoechst 33342 is a potential carcinogen. Proper PPE should be used while handling.
17. Wash the secondary antibody solution with PBS-Tw 3×5 min.
18. Fix samples with PFA for 15 minutes at room temperature.
19. Wash the fixative with PBS-Tw and equilibrate samples within 60% glycerol. Mount explants on microscope slides with nail polish and 60% glycerol for imaging. For representative immunostaining results please see Figure 3.