Sectioning & Staining 

Sectioning Procedure:

1. Cut straight across the cerebellum so the brain can stay level and upright.

2. Apply 1% Ethanol to the frozen stage as the base.

3. When the Ethanol is halfway frozen, add a layer of 30% sucrose to the base.

4. When the sucrose is halfway frozen, place the brain into/onto the stage with the dorsal side facing towards the blade. 

5. Add more sucrose around the tissue and allow it to freeze.

6. Manually lower or raise the stage so that the tissue is below where the blade will run.

7. Place the blade in its designated space and tighten the screws to hold it the blade into place.

8. Obtain a brush to handle the tissue and a sectioned dish full of distilled H2O to place tissue.

9. Begin cutting tissue into sections of the desired thickness (50 microns) and fully extend cutting arm for the automatic adjustments. Place each section of tissue into the sectioned dish filled with distilled water.

10. When cutting is all done, remove the blade, wash it with soap and water. Dry the blade and coat it in oil, store it in its box until next use.

11. For mounting, place one section in a dish of water, let it float to the top and then place the glass slide under the tissue in the water, use the brush to get the tissue onto the slide. Repeat for the other pieces of tissue.

12. When done mounting the tissue to the glass slide, allow the tissue to dry before staining.

Staining Procedure:

1. Fill up 4 beakers with one of these different solutions: water, 70% alcohol, 100% alcohol, and a clear safe.

2. Immerse the tissue glass slides into the beaker filled with a water solution.

3. Immerse the tissue glass slides into the beaker filled with a 70% Alcohol solution.

4. Immerse the tissue glass slides into the beaker filled with a 100% Alcohol solution.

5. Immerse the tissue glass slides into the beaker filled with clear safe solution.

6. Take the glass slide from the clear safe solution and immerse it into the beaker filled with a 100% Alcohol solution.

7. Take the glass slide from the 100% Alcohol solution and immerse it into the beaker filled with a 70% Alcohol solution.

8. Take the glass slide from the 70% Alcohol solution and immerse it into the beaker filled with a water solution.

9. Take the tissue slides from the water solution and immerse it into the beaker filled with the nissl stain solution. Let it sit for a few minutes.

10. Fill another beaker with 10% Acid ETOH.

11. Take the tissue slides from the stain and immerse it in the beaker with the water solution, then spill out the solution in the hazardus waste bucket while holding the stain slides in place. Follow by refilling the beaker with the water solution. 

12. Take the tissue slides from the water solution and immerse it in the beaker with the 10% Acid ETOH solution, then spill out the solution in the hazardus waste bucket while holding the stain slides in place. Follow by refilling the beaker with the 10% Acid ETOH solution. Spill the beaker of the water solution into the hazardus waste bucket.

13. Take the tissue slides from the 10% Acid ETOH solution and place them into the beaker with the 70% Alcohol solution, followed by immersing the slides in the 100% Alcohol solution. Leave the slides in the 100% solution for a couple minutes. Spill the beaker of 10% Acid EtOH solution into the hazardus waste bucket.

14. Take the tissue slides from the 100 % Alcohol solution and place them into the beaker with the clear safe solution. Spill out the beaker of 100% Alcohol solution into the hazardus waste bucket.

15. Place a swab into Toluene Solution (SP15-100) and separate the cover slips for the slides. 

16. Take one slide from the clear safe solution and use the swab dipped in the Toluene Solution to spread a line across the top of the slide. Place the cover slip from the top of the slide and let it lay on its own. This will allow the toluene solution to spread over the whole tissue on the slide. Repeat with the other slides.