Sectioning and Staining Procedure

Sectioning Procedure:

1. First Open the valve to CO2 tank and cool stage until completely frozen

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

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

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

5. When Sucrose is halfway frozen, place the brain so that the dorsal side will face toward the blade.

6. Add more sucrose around the tissue and allow to freeze.

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

8. Place the blade in its designated space and tighten the screws to hold the blade in place.

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

10. Begin Cutting Tissue into sections of the desired thickness (50 microns) and fully extend the cutting arm for automatic adjustments.

11. When done cutting, remove the blade, wash with soap and water, then dry. Coat the blade in oil and store until the next use.

12. For mounting, place one section in a dish of water, float to the top, and place slide under tissue in water.

13. Using a brush, guide the tissue to its spot on the slide.

14. When done mounting, allow your tissue to dry before staining. 

Staining Procedure:

1. You need 6 containers. Five in the front and the sixth one in the back for later use. 

2. Fill the first one with distilled water, the second with 70% alcohol, the third with absolute alcohol, the fourth with clear safe and the fifth with Cresyl Violet stain. 

3. Take slides with the backside facing outwards 

4. Add all slides to the distilled water into the correct slots. 

5. Remove the slides from the distilled water and transfer them to container with the 70% alcohol 

6. Soak them in 70% alcohol for 2-4 mins to properly dehydrate the tissue. 

7. Move them into the absolute alcohol for 5 mins to complete dehydration. 

8. Then into clear safe. If you see smoke, this means that the dehydration process has not been complete, so you must move slides back into the alcohol for several minutes before adding them back to the clear safe. 

9. Leave the slides in the clear safe for 5 minutes or until the tissue becomes transparent. 

10. Move the slides back into the absolute alcohol for 4 minutes

11. Transfer slides into the 70% alcohol another 4 mins

12. And then into the water for an additional 2 minutes. 

13. Finally add the slides to the stains for at least 10 mins. 

14. While the slides are soaking in the stain, prepare the sixth container by filing it with acid alcohol.

15. Place the sixth container with the acid alcohol between the containers containing distilled water and the 70% alcohol. 

16. After 10 minutes, transfer the slides out of the stain to the water. As water changes color, dump out water and refill it. 

17. Then move the slides to the acid alcohol, where it should turn pink. Dump out the pink liquid and refill it with acid alcohol. 

18. Add the slides to the 70% alcohol, dump out colored liquid and refill it. 

19. Move into abs al for several mins, do not dump, slides should not be losing a lot of dye at this point in the process. 

20. Then finally add the slides to the clear safe, where they should not be losing anymore dye. 

21. Prepare cover slips, do this without gloves to more easily separate the cover slips. 

22. Get as many coverslips as slides that you have prepared. 

23. Put gloves back on. Use a one-sided Q-Tip and dip into Permont. 

24. Remove the slides from the clear safe with the brains facing upwards, and hold it at an angle facing away from you

25. Then apply Permont to the bottom edge facing away from you and place the coverslip perpendicular to the slide about halfway down the coverslip and move it at an up-increasing angle until it is parallel with the slide.

26. Tap the slide on each side. 

27. Repeat with the remaining slides and set them aside to dry.