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.
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.
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.
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.
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.
At this point the sections will look blue.
* 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.
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. | Deparaffinize and rehydrate to water |
| 2. | Clear-Rite 3 3 times for 3 min each |
| 3. | 100% ethanol 3 times for 1 min each |
| 4. | 80% ethanol 30 sec |
| 5. | 50% ethanol 30 sec |
| 6. | 30% ethanol 30 sec |
| 7. | Rinse is slow running water 30 sec |
| 8. | Carbol-fuschin 30 min |
| 9. | Rinse in distilled water |
| 10. | 1% glacial acetic acid in 70% ethanol about 1.5 min, depending on tissue* |
| 11. | Rinse in running tap water 10 min |
| 12. | Methylene blue short dip |
| 13. | Rinse in running tap water 5 min |
| 14. | 95% ethanol 6 dips |
| 15. | 100% ethanol 2 min |
| 16. | 100% ethanol 5 min |
| 17. | xylene 3 min (repeat 3 times) |
| Carbol-fuschin | |
| Basic fuschin | 1 g |
| Melted phenol | 5 g |
| 100% ethanol | 10 ml |
| Distilled water | 100 ml |
| Filter before use, can be re-used | |
| Methylene Blue | |
| Methylene blue | 0.14 g |
| 95% ethanol | 10.0 ml |
| Tap water | 90 ml |
Dissolve stain in ethanol, then add water. Discard after use.