Zebrafish Scale Regeneration In Toto and Ex Vivo Culture of Scales

The University Animal Scientific Unit (ASU) is responsible for zebrafish care under the guidance of zebrafish husbandry guidelines. All procedures, including scale harvesting, live bone staining, and live imaging, were approved and performed under a UK Home Office Project License (PP4700996). For this manuscript, young adult transgenic zebrafish from the sp7:mCherry [Tg(osterix:mCherry-NTR)^pd46] line was used¹⁷. The fish included both males and females of 4 months old.

1. In vivo scale regeneration

1. Before starting the scale regeneration experiment, transfer zebrafish from their main tanks into individual tanks (see Table of Materials) and return to these individual tanks following imaging throughout the experiment, as shown in Figure 2. This is to ensure that all regenerating scales are those harvested for the experiment; group housing can lead to sporadic loss of scales due to injuries caused by other fish.
   NOTE: The fish are individually placed to avoid fighting and subsequent scale loss between fish.
2. Record information for each experiment fish (i.e., genotype, age, and sex) according to the experiment design. Use a camera (smartphone cameras work fine) to take an image of each fish next to a ruler to record fish size/length and health condition.
3. On the day of the experiment, anesthetize the zebrafish using 0.05% (v/v) tricaine methane sulfonate (MS222, see Table of Materials) by immersion, then place it on a Petri dish with a damp tissue to avoid movement of the fish during imaging or harvesting.
4. Place the fish on its side (typically left flank is used here for consistency). Perform flank imaging using a stereomicroscope with appropriate magnification (typically, 2x, 4x, and/or 6.3x and 8x were used for the present study to capture both the overall regenerating area and the zoomed-in area for detailed observation).
   NOTE: The flank imaging timepoints depend on the experimental purpose and choice of transgenic reporters. For example, to track osteoblasts during scale regeneration, the suggested timepoints are Day 0 (ontogenetic), 2 days post-harvest (dph), 4 dph, 7 dph, 10 dph, 14 dph, 21 dph. To reduce effects from repeated anesthesia, keep imaging timepoints to the fewest required for the experiment.
5. Harvest scales using fine tweezers and forceps under the stereomicroscope. Transfer the harvested scales into collection tubes for scale staining later.
   NOTE: The maximum number of scales that can be harvested will depend on local ethical approvals that are in place. We typically harvest around 20 to 30 scales and collect them into individual collection tubes for scale staining later, such as ALP, von Kossa¹⁴.
6. Collect the scales into either a 1.5 mL or 2 mL collection tube. This depends on the staining technique used.
   1. For both ALP and von Kossa staining, transfer the scales into collection tubes containing deionized water, whereas for TRAP staining or immunohistochemistry, transfer the scales into collection tubes containing 4% PFA (fixing solution).
7. Optional: Before transferring scales into tubes, capture images of individual harvested scales if required.
   NOTE: Suggested scale harvest timepoints are day 0 (ontogenetic), day 10, and day 21 (regenerated scales).

2. Live bone staining

NOTE: Live skeletal staining can be performed either when experimental fish are not transgenic for fluorescent reporters or when carrying single-color transgenic reporters. Live staining can be used to complement transgenics; for example, one might use an osteoblast reporter such as sp7/osx in one color (e.g., GFP), then stain bone in red with Alizarin red (AR). AR and calcein green can be used for the same fish together; in this scenario, AR is used to label original or ontogenetic bone by binding to the mineralized matrix of aged bone, and calcein binds to calcium ions that are abundant in newly formed bone. For example, when staining an ontogenetic scale, AR can be visualized, but calcein green may be absent or minimal since ontogenetic scales have low levels of new bone formation.

1. Perform live Alizarin Red (AR) staining.
   1. Prepare a bottle of 2x AR staining solution by mixing 0.5% (w/v) stock Alizarin solution (10 mL) with 10 mL of 1 M stock HEPES (final concentration, 10 mM) (see Table of Materials).
   2. Top up to 1 L with 1x Danieau solution (see Table of Materials). The solution must be kept in dark or wrapped with foil.
   3. On the day of the experiment, bring 500 mL of 2x AR solution to the zebrafish facility, and add 500 mL of system water (from the aquarium) to make 1x working AR solution.
   4. Transfer fish from their tanks into a 1x working AR staining solution and leave for 15-20 min. Wash off the excess stain on the fish with 15 min 'washing' in the system water.
2. Perform live Calcein Green staining.
   1. Prepare a bottle of 2x Calcein Green staining solution by adding 45 mg of Calcein powder into 900 mL of 1x Danieau solution (see Table of Materials). Use a magnetic stirrer to allow the powder to dissolve completely.
   2. Adjust the pH to 8. This is the 2x stock Calcein Green solution. The solution can be stored in the dark/wrapped with foil for up to 1 month.
   3. On the day of the experiment, bring 500 mL of 2x Calcein Green solution to the zebrafish facility, and add 500 mL of system water to make 1x working Calcein Green solution. It is recommended to use freshly made Calcein staining solution.
   4. Transfer fish from their tanks into 1x working Calcein Green staining solution and leave for 1-2 h. Depending on the size of the fish, a longer staining time may be required.
   5. Wash off the excess stain on the fish by 15 min 'washing' in the system water.
      NOTE: Image the Calcein Green stained fish as soon as possible as this stain will fade in a few hours.

3. Alkaline phosphatase (ALP) staining on scales post-harvest

1. Prepare the ALP staining solution by mixing 100 mM Tris (pH 9.5 with HCl) with 100 mM NaCl and 50 mM MgCl₂.
2. Use commercially available NBT/BCIP stock solution (see Table of Materials).
3. Transfer the scales into tubes containing deionized water.
4. Rinse the scales in ALP staining solution for 5 min.
5. Meanwhile, prepare a working staining solution by adding 200 µL of the NBT/BCIP solution to 10 mL of ALP staining buffer. This needs to be freshly prepared at the time of staining.
6. Stain the scales with the working ALP staining solution for 15 min.
7. Stop the reaction by rinsing the scales with deionized water.

4. Von KOSSA staining on scales post-harvest

1. Prepare 5% (w/v) silver nitrate solution.
2. Prepare 5% (w/v) sodium thiosulfate solution.
3. Transfer harvested scales into tubes containing deionized water.
4. Incubate the scales in the silver nitrate solution for 40 min under strong light.
   NOTE: Wear PPE (personal protective equipment) as silver nitrate leaves a stain that is difficult to remove.
5. Wash the scales twice for 5 min with deionized water.
   NOTE: Silver nitrate waste must be collected and disposed of according to local guidelines.
6. Incubate the scales in sodium thiosulfate for 5 min.
7. Wash the scales twice for 5 min with deionized water.

5. Colorimetric TRAP staining

NOTE: For the detailed procedure, please refer to previously published work¹⁸.

1. Prepare the staining solutions.
   1. Prepare 10 mg/mL Naphthol AS-MX phosphate solution.
      1. Weigh 5 mg Naphthol AS-MX phosphate (see Table of Materials).
      2. In the fume hood, prepare a tube with 0.5 mL of N, N'-dimethylformamide and dissolve 5 mg of Naphthol AS-MX phosphate in 0.5 mL of N, N'-dimethylformamide to make a stock solution of 10 mg/mL of Naphthol AS-MX phosphate (shake gently).
   2. Prepare 1.6 mM of Fast Red Violet LB stock solution in 50 mM sodium tartrate.
      1. Prepare 50 mL of 0.1 M sodium acetate at pH 5 (add 0.41015 g of sodium acetate in 50 mL of deionized water and adjust the pH to 5 with 100% acetic acid).
      2. Dissolve 0.575 g sodium L-tartrate dibasic dihydrate (see Table of Materials) in 50 mL of 0.1 M sodium acetate buffer (pH = 5).
      3. Add 30 mg of fast red violet LB salt.
   3. Prepare PBS-0.1Tx (dissolve 500 µL of Triton-X in 500 mL of PBS).
      1. Prepare PBS-0.1Tw (dissolve 500 µL of Tween-20 in 500 mL of PBS).
2. Prepare a working TRAP solution.
   1. Mix both stock solutions together in the fume hood (0.5 mL of 10 mg/mL Naphthol AS-MX phosphate solution and 50 mL of 1.6 mM Fast Red Violet LB stock solution in 50 mM sodium tartrate).
      NOTE: The working solution can now be used outside the fume hood. It can be stored in the fridge (wrapped in foil) for up to a month. Store in separate containers from antibodies or other reagents sensitive to fixatives. Keep away from light as much as possible.
   2. Optional: prepare bleaching solution (on post-fixed samples that were TRAP stained) in PBS and 0.1% Tween-20 (PBS-0.1%Tw).
   3. Prepare a final concentration mix of 0.5% potassium hydroxide (KOH) (from 10% stock solution (w/v)) and 3% hydrogen peroxide (H₂O₂) (33% stock solution, stored in the fridge).
3. Perform sample fixation.
   1. Fix scales in 4% PFA in 1x PBS rocking or rotating for 40 min at room temperature (RT).
   2. Remove the 4% PFA and wash with PBS-0.1Tx at least 3 times for 5 min each wash.
   3. Directly move to TRAP staining (the best approach) or store in PBS-0.1 Tx overnight.
4. Perform TRAP staining of the scales.
   1. Remove the PBS-0.1 Tx solution and add the colorimetric TRAP working solution (step 5.2.1) to cover the samples fully (i.e., 1 mL in a 1.5 mL microcentrifuge tube).
   2. Incubate for 1-2 h at RT, rocking in the dark.
   3. Remove the staining solution and wash at least 3 times with PBS-0.1 Tw for 5 min each wash.
   4. Post-fix the samples in 4% PFA for 30 min at RT, gently rocking in the dark.
   5. Remove the PFA solution by washing 3 times with PBS-0.1 Tw for 5 min each step.
      NOTE: The samples can be stored (in storage solutions/mounted on microscope slides) at 4 °C in the dark indefinitely.

6. Mounting of stained scales

1. Prepare the mounting medium by adding 2.4 g Mowiol 4-88 crystals to 6 g of glycerol (see Table of Materials). Any preferred mounting media can be used.
2. Add 6 mL of deionized water and leave on a magnetic stirrer for an hour.
3. Add 12 mL of 0.2 M Tris (pH 8.5) and incubate at approximately 53 ° C until the Mowiol crystals fully dissolve.
4. Clarify the solution by centrifuging at 2000 x g for 20 min at room temperature.
5. Transfer the solution to a storage container. The mounting medium can be kept in the freezer for about 12 months. The defrosted Mowiol solution is stable for up to 1 month.
6. Mount the scales on a microscope slide in mounting the medium, cover them with a coverslip and view them under a microscope.

7. Ex vivo culture of scales

NOTE: This step is adapted from de Vrieze et al.¹².

1. Before harvesting the scales in a sterile environment, prepare the osteogenic mediumand the culture plate.
   1. Osteogenic Culture Medium (OCM): In a sterile centrifuge tube, make 15 mL OCM in standard cell culture medium (high glucose, no glutamine, no phenol red) at the final concentrations of 1% bovine calf serum, 1% (200 mM L-alanyl-L-glutamine dipeptide in 0.85% NaCl), 1% antibiotic/antimycotic solution (100x) solution, 4 mM CaCl₂, 10 mM β-Glycerophosphate, 1 nM sodium pyruvate and 1 % Amphotericin B (optional) (see Table of Materials).
   2. Pour the OCM into a sterile reagent reservoir, and using a multichannel pipette, add 100 µL to each well of the 96-well plate.
   3. Seal the 96-well plate and store the plate. OCM can be stored in the refrigerator for a short period (overnight to 1 week) and kept at 28 °C before use.
2. Perform scale harvesting and PBS washing.
   1. Harvest the scales using sterile tweezers/forceps and deposit them in a Petri dish containing a sterile PBS solution.
3. Perform plate setup.
   1. Transfer each scale from the Petri dish to each well on the 96-well plate with 100 µL of OCM (warmed to 28 °C) using sterile forceps.
   2. Once all scales are transferred to the plate, bring the plate under a stereomicroscope. Using a fine, sterile pipette tip to gently move each scale to the bottom of the well and keep them away from the side of the wells to avoid light reflection during image acquisition.
      NOTE: This step is required only if performing live imaging.
   3. Carefully transfer the plate to the incubator or Live Imaging System (LIS, see Table of Materials) (set at 28 °C, 5% CO₂), where the scales can be cultured for up to 7 days.
4. Refresh OCM (optional).
   1. Warm the OCM that was stored in the fridge to 28 ˚C.
   2. Carefully transfer the 96-well plate to a sterile environment. Using a multichannel pipette, aspirate 50 µL from each well. For this, press the tips against the walls and do not dip them until the bottom to avoid aspirating/moving/removing the scales. Use new tips for different conditions.
   3. Pour the OCM into a sterile reagent reservoir, and using the multichannel pipette add 60 µL of OCM to each well.
   4. Place the plate under the stereomicroscope and check for misplaced scales (too close to the walls, floating, or in a vertical position within the media). Reposition them with a fine, sterile pipette tip if needed.
   5. Place the plate back in the incubator/LIS.
      NOTE: Do not refresh the medium if the experiment involves circadian rhythm (CR), as serum shock can affect the biological clock. Note if studying CR notes that light conditions also reset the clock.
5. Perform post-culture scale staining and imaging (optional).
   1. After 6 days of culture, fix them and direct them for appropriate staining such as von Kossa and ALP mentioned above.
   2. Alternatively, if the experimental fish have a fluorescent tag, put the plate into a live imaging system.
      NOTE: If using a live imaging system, ensure that it is set to a temperature and CO₂ level appropriate for the experiment. Typically, in this study, 28 °C and 5% CO₂ were used. Select appropriate filters and imaging timepoints (here, an interval of 2 or 4 h, which is sufficient to follow osteoblast migration, was used) and low magnification objectives (i.e., 4x) to ensure even imaging of the entire wells. This is because the scales tend to move inside the wells, particularly during media changes.