Recipes and Protocols

• 95% ethanol 300 ml
• formalin 200 ml
• glacial acetic acid 100 ml
• distilled water 300 ml

• 95% ethanol30 mL
• formalin10 mL
• glacial acetic acid 2 mL
• distilled water58 mL
• 100 mL total

Solution A 2.62 % NaH₂PO₄

Solution B 2.52 % NaOH

Mix 83 ml Solution A with 17 ml Solution B.

Adjust pH to 7.4 with 10 N NaOH or concentrated HCl.

For transmission electron microscopy of fish tissues, we generally prepare 4% glutaraldehyde in Millonig's buffer. Fix small pieces of tissue at room temperature for 2 h, then hold overnight at 4 ⁰C. Transfer to Millonig's buffer without glutaraldehyde.

• tryptone 0.5 g
• yeast extract 0.5 g
• sodium acetate 0.2 g
• beef extract 0.2 g
• agar 11.0 g

• distilled water 1 liter
• autoclave and pour into sterile petri dishes

This stain is useful for examining protozoa, bacteria and cell morphology in tissue imprints and blood smears. Pre-made commercial products, such as Diff-Quik (Dade Behring AG, Newark, DE) http://www.dadebehring.com/ are nearly equivalent.

Add 1 g Leishman's stain to 500 ml absolute methanol and filter.

Add 1 g powder to 66 ml to glycerol and heat at 60 ⁰C for one hour. Then add 66 ml of absolute methanol and filter. Store at 40C.

• Solution A - 31.20 g NaH₂PO₄ , 2 H₂0 in 1 L distilled water
• Solution B - 53.65 g Na₂HPO₄ , 7 H₂0 in 1 L distilled water

To prepare phosphate buffer, mix 73.5 ml solution A with 26.5 ml solution B. Then add 100 ml distilled water. Add 3.5 ml stock solution to 50 ml phosphate buffer (pH 6.4). Make fresh for each use.

• Prepare smear or imprint and allow to dry for about 1/2 hour.
• Fix in methanol for 2-5 min.
• Stain in Leishman's for 2-3 min.
• Stain in Giemsa for 10-12 min.
• Rinse in distilled water for about 1 min.
• Air dry.

• Fish should still be alive just prior to fixation.
• Ice the fish to kill them. Cut off the tail behind the anus. Slit open the body cavity along the belly.
• Use 15 mL fixative per 1-2 fish in a small vial.
• To ensure uniform and complete fixation, fix for three days on rotor, or other agitating device.
• Store fish in Dietrich’s fixative at room temperature

Adult zebrafish are often difficult to section without first softening their skin and scales. The day before processing, transfer the fixed fish to 5% trichloroacetic acid (TCA) in Dietrich’s fixative, and place on rotor overnight. TCA will soften the scales and skin, presumably by cleaving the keratin proteins. In addition, TCA will decalcify, softening the bones. Unfortunately, TCA is a fairly strong acid and will corrode the tissue processor. The fish must be thoroughly rinsed to remove any TCA before the fish can be processed.

Transfer fish to 5% TCA in Dietrich’s fixative. Place on rotor overnight. The next day, replace TCA-containing Dietrich’s with 70% ethanol and place on rotor for 10 minutes. Repeat 2xs for a total rinse time of 30 minutes. Replace last rinse with fresh 70% ethanol. The fish are then bisected along their length before they are processed. Using a fresh razor blade, make the cut parallel to and on the left side of the spinal cord. The bisected fish are now ready to load into cassettes for processing.

• 70% ethanol 10 minutes
• 70% ethanol 40 minutes
• 80% ethanol 1 hour, 10 minutes
• 95% ethanol 2 X 1 hour, 10 minutes
• 100% ethanol 2 X 1 hour, 10 minutes
• xylene 1 hour, 30 minutes
• Clear-Rite 3 (from Richard-Allen Scientific) 1 hour, 30 minutes
• Paraplast/Paraplast Plus* (50:50) 2 X 1 hour
• Paraplast 1 hour
• Paraplast 45 minutes
• Embed in TissuePrep-2 Embedding Media (Fisher Scientific), placing cut sides of both halves of the fish down.
• *Paraplast Plus contains DMSO. Care should be exercised when handling used Paraplast Plus as the DMSO can carry residual xylene across the skin.

• 70% ethanol 10 minutes
• 70% ethanol 50 minutes
• 80% ethanol 50 minutes
• 95% ethanol 2 X 50 minutes
• 100% ethanol 2 X 50 minutes
• xylene 1 hour, 10 minutes
• Clear-Rite 3 (from Richard-Allen Scientific) 1 hour, 10 minutes
• Paraplast/Paraplast Plus* (50:50) 2 X 1 hour
• Paraplast 1 hour
• Paraplast 45 minutes
• Embed in TissuePrep-2 Embedding Media (Fisher Scientific), placing cut sides of both halves of the fish down.
• *Paraplast Plus contains DMSO. Care should be exercised when handling used Paraplast Plus as the DMSO can carry residual xylene across the skin.

• Fish younger than 3 weeks can be pre-embedded in Histogel, available from Fisher Scientific, Richard Allan Scientific or Lab Storage Systems.
• 70% ethanol 2 X 20 minutes
• 80% ethanol 30 minutes
• 95% ethanol 2 X 30 minutes
• 100% ethanol 2 X 30 minutes
• xylene 40 minutes
• Clear-Rite 3 (from Richard-Allen Scientific) 40 minutes
• Paraplast/Paraplast Plus* (50:50) 2 X 40 minutes
• Paraplast 40 minutes
• Paraplast 15 minutes

If you want serial sections, embed in Paraplast as this paraffin provides for better ribboning.

Cut 7 µm sections with a high-profile disposable blade (AccuEdge blades – made by Sakura-Finetek, sold through VWR – are reportedly the best) . If the large fish are difficult to cut, soak the block face in ice-cold Mollifex. Usually one to two hours is sufficient but an overnight soak is fine for stubborn samples.

Sections of gills, spinal cord and internal organs will be needed for proper pathological evaluation. Because the fish has been cut to one side of the spinal cord, you will be cutting toward the midline on one half of the fish, and away from the midline on the other. With any luck, you will need to collect only one ribbon containing gills (one bisected half) and spinal cord (the other bisected half), along with a nice sampling of internal organs. Samples of the skin or fins may also be requested, depending on the reported symptoms.

Mollifex: 54 mL 95% ethanol, 10 mL glycerol, 36 mL water

Store fixative at room temperature.

The following is a staining protocol perfected by Karen Larison at ZIRC that we routinely use for zebrafish histopathology. Any water stage is a good stopping point if you had to do something part way through the procedure. Keep the slides wet throughout the procedure.

• Clear-Rite 3 3X 3min
• 100% ethanol (absolute) 3X 1 min
• 80 % ethanol 30 sec
• 50 % ethanol 30 sec
• 30 % ethanol 30 sec
• Rinse in slow running water 30 sec
• Harris' Hematoxylin * 4.5 min
• Rinse in slow running distilled water till stain rinsed out ~4 min
• Decolorize in 0.1% Acid alcohol** 45 sec
• Rinse in distilled water 15 sec
• Lithium carbonate neutralizing station** * 8 dips

At this point the sections will look blue.

• Rinse in slow running water minimum of 5 min
• 95 % ethanol 30 sec
• 95% ethanol 30 sec
• eosin Y - phloxine B solution**** 30 sec
• 100 % isopropanol***** 6X 15 sec
• xylene 3X 15 sec
• Coverslip with permount.

* Filter Hematoxylin before each use. Make fresh batch every 2-3 months.
** 200 ul concentrated HCl in 200 mL 70% ethanol
*** Water and lithium carbonate (approx 2 tablespoons, supersaturated)
**** Make fresh eosin Y- phloxine B solutions from stock every 1-2 weeks.
***** Use fresh isopropanol for the 6th wash. Discard the 1st (and pinkest) isopropyl. Rinse in isopropyl waste container after each staining session. Store other used rinses in bottles 1-5 (pinkest to clearest) for future use.

• Hematoxylin 2.5 g
• Absolute alcohol 25 ml
• Potassium alum 50 g
• Distilled Water 500 mL
• Sodium iodate 0.5 g
• glacial acetic acid 20.0 mL

The hematoxylin is dissolved in the absolute alcohol, and then added to the alum, which has previously been dissolved in the warm distilled water in a 2-liter flask. The mixture is rapidly brought to a boil and sodium iodate is then slowly and carefully added. The stain is rapidly cooled by plunging the flask into cold water or into a sink containing chipped ice. When the solution is cold, the acetic acid is added, and the stain is ready for immediate use.

• 1% eosin Y (aq) 25.0 mL
• 1% phloxine B (aq) 2.5 mL
• 95% ethanol 195.0 mL
• glacial acetic acid 1.0 mL

Dissolve stain in ethanol, then add water. Discard after use.

• Step 10. This step is subjective. The goal is to decolorize tissues while bacteria retain the stain.