Histology Protocol

Fixation

  1. 1.5 ml FSW (0.2 µm filtered seawater) in a 15 ml tube (for small fragments).
  2. Cut the sample into approximately 2 cm fragments and place them in the tube.
  3. Leave the fragments in the tube with the FSW for 10 min before adding 1M MgCl₂.
  4. Add 0.3 ml of 1M MgCl₂ to a single tube, and cover the plate with aluminum foil for 10 min.
  5. Remove the FSW from the tube.
  6. Add 4% formaldehyde solution into the tube.
    Second option: leave the 1.8 ml of FSW and 1M MgCl₂ solution in the tube and add 0.5 ml of 16% formaldehyde to the tube for a final concentration of 4%.
  7. Leave at 4°C overnight or 1–2 hours at room temperature.
  8. Wash ×3 with sterile PBS solution.
  9. For extended storage, store in 70% sterile ethanol, or leave in PBS for short storage of 2–3 days.

Paraffin Embedded Tissue

  1. Wash ×3 with PBS at room temperature, 15 min each.
  2. For scleractinian: decalcify skeleton with 1:1 formic acid (50% in DDW) & sodium citrate 20% (20% in DDW).
    Transfer the fixed sample into 50 ml tubes and add the decalcification solution (10 ml in each tube; for samples >3 cm add 15 ml) for 4–5 hours/overnight.
  3. Wash ×3 with PBS at room temperature, 15 min each.
  4. Dehydrate tissue through graded ethanol (with PBS): 50%, 75%, 90%, 100% (15 min each).
    If tissue >2 mm thick, extend 100% step to 30 min.
  5. Clear tissue through graded xylene or butanol (with ethanol):
    • 50% butanol:ethanol, 30 min
    • 75% butanol:ethanol, 30 min (samples can be stored overnight at 4°C)
    • 90% butanol:ethanol, 30 min
    • 100% butanol:ethanol, 30 min
  6. Add 15 ml paraffin to 50 ml tube and melt in incubator at 58–60°C (~2 hours).
  7. Remove butanol and insert tissue into tube (#1) with melted paraffin. Incubate at 58–60°C for 1 hour.
    Tip: transfer tissue quickly using a plastic spatula.
  8. Transfer tissue into tube (#2) with melted paraffin. Incubate overnight at 58–60°C.
  9. Transfer tissue into tube (#3) with melted paraffin. Incubate 2 hours at 58–60°C.
  10. Place tissue in correct position on mold.
  11. Pour melted paraffin until tissue is fully covered.
  12. Chill on ice for a few hours, then transfer to 4°C overnight.

Storage: Save paraffin blocks at room temperature. In hot environments, store blocks in a box at 4°C. Before sectioning, chill blocks on ice or in ice-cold water.

Tissue Sectioning (Microtome)

Equipment: Knives, acetone, brush, DDW, pencil, slides, slide boxes, scalpel, tweezers, ice.

  1. Remove block from mold and mark tissue area with scalpel.
  2. Peel edges until tissue square protrudes.
  3. Place block on ice for a few minutes.
  4. Clean knife and surface with acetone.
  5. Mount block on microtome and adjust knife.
  6. Slice sections (10 µm each).

Stains

Equipment: Large beaker with DDW, small beaker for waste, pink tubes with ethanol, 100% ethanol, glass pipette, stains kit.

  1. Place 1 drop of hematoxylin on each tissue slice (1 min).
  2. Preliminary wash with pipette into waste tank.
  3. Second wash: 10 sec in DDW beaker.
  4. Place 1 drop of bluing reagent (15 sec).
  5. Preliminary wash with pipette.
  6. Second wash in DDW beaker.
  7. Dip slides in pink tube #1 (100% ethanol, 10 sec).
  8. Dip slides in pink tube #2 (100% ethanol, 10 sec).
  9. Place 1 drop of eosin on each tissue slice (2–3 min).
  10. Wash with 100% ethanol (10 sec, pipette).
  11. Dehydrate with 100% ethanol (pink tubes), 1 min ×3.

Coverslip

Equipment: Scotch super glue, cover glasses, 80% glycerol.

  1. Place 2 drops of 80% glycerol on slide center (enough to cover samples).
  2. Place 4 small drops on cover glass corners and immediately mount.

Notes:

  • Do not dehydrate sample before glycerol placement.
  • DPX glue can be used instead (ensure no dehydration before gluing).
  • DPX gluing must be done in a hood.

Reference

Written on March 9, 2022
Citation:
Gametogenesis: Oceans 2024, 5(4), 758–769; https://doi.org/10.3390/oceans5040043

Gametogenesis assessed using histological sections of corals throughout the year. Coral fragments were placed in 15 mL tubes with 10 mL filtered seawater and 2 mL of 1M MgCl₂ for 15 min to prevent contraction. Samples were fixed in 4% formaldehyde for 24 h, rinsed in PBS, and preserved in 70% ethanol. Decalcification was performed with 1:1 formic acid (50% in DDW) and sodium citrate (20% in DDW) [Rinkevich & Loya, 1979]. Tissue was dehydrated in graded ethanol, cleared in butanol, and embedded in paraffin. Sections (10 µm) were stained with hematoxylin and eosin, then analyzed under a Nikon ECLIPSE Ti2 inverted microscope. Gonads per polyp were counted, oocyte diameters measured using Nikon Nis-Elements software (v5.02).

Written on December 6, 2020