Mammary Gland Whole Mount Processing and Staining

Note: Steps 1 - 22 are performed manually using stainless steel staining dishes.

1. Remove the mammary gland from the animal and spread onto a clean 75x50mm glass microsope slide (Fisher Scientific, # 12-553-5B) as quickly as possible. Fix in 10% neutral buffered formalin or methacarn for 18 - 24hrs. If you are not processing the whole mounts immediately, pour off the fixative into a suitable container for disposal and immerse the whole mounts in 70% ETOH. Cover the staining dish with parafilm and place the cover on top. 

2. Rinse formalin fixed tissue in running tap water for 15 min. (Do not rinse methacarn fixed tissue).

3. Dehydrate in 95% ETOH for 1 hr.

4. Dehydrate in 100% ETOH for 1hr.

5. Clear in xylene for 2 hrs.

6. Rinse in 100% ETOH for 1 hr.

7. Hydrate in 95% ETOH for 1 hr.

8. Rinse in running tap water for 15 min.

9. Stain in Alum Carmine until stain has infiltrated tissue 2 - 5 days depending upon the thickness of the specimen and the freshness of the stain.


10. Rinse in running tap water for 15 min.

11. Dehydrate in 95% ETOH for 1 hr

12. Dehydrate in 100% ETOH for 1 hr

13. Clear in xylene for 2 hrs.

14. Remove whole mounts from xylene one at a time and drain excess xylene. (Please note: If you want to coverslip the whole mounts instead of bagging them now is the time to do so.)

15. Place each whole mount into a 4"x 6" Kapak™ (Fisher Scientific, # 01-812-25D) heat seal bag and label the outside corner portion of the bag with the study and animal # or designated code. To view examples of steps 15 - 20 click here.

16. Fill the bag with 20ml of methyl salicylate (Sigma Chemical, # M6752) and remove as much of the air as possible. Please note that methyl salicylate is highly aromatic and will act as an intoxicant in large quantities. It is recommended that this compound be used under a chemical fume hood with a NIOSH approved air purifier that has organic filters. In addition, double gloves should be worn to avoid exposure and the outer pair should be changed frequently. 

17. Seal the top portion of the bag using the Kapak™ heat sealer (Fisher Scientific, # 01-812-15), crimp and hold for 5 sec. Air bubbles may form as a result of the residual xylene mixing with the methyl salicylate. For this reason, set the whole mount aside until the next day.

18. Remove residual air bubbles from the whole mount by laying the bag on a flat surface, whole mount side up. Place index finger on the bag press and squeeze air bubbles to the side forcing them out from under the tissue. Hold the bag vertical and tap the bottom of the bag to remove any air.

19. The heat sealer has been modified (the middle plastic bar has been removed leaving a 4" slit in the top of the unit which allows the bags to be inserted vertically from the top down. Insert the bag in the heat sealer starting from the top and going down. The bottom edge of the bag should be even with the bottom of the sealer. While holding the top of the bag with the right hand place the back of the left hand press the whole mount up against the sealer, this will force the excess methyl salicylate up. While the bag is pressed against the sealer with back of left hand use the right hand to crimp the bag in the middle (hold for 5 sec.) Release, and pull the bag out through the bottom of the heat sealer.

20. Using scissors, cut along the crimp line being careful not to nick the bag and release the contents. The top portion of the bag (does not contain the whole mount) will be retained and the methyl salicylate will be recycled.

21. Photograph the whole mounts for archival purposes (refer to W.M. photo procedure).

22. Excise lesions from the whole mounts one at a time an place into a tissue processing cassette. The cassettes are then loaded into an automatic tissue processor. 

23. Clear in Toluene for 1hr.

24. Clear in Toluene for 1hr.

25. Infiltrate with molten paraffin (paraplast+) for 1hr.

26. Infiltrate with molten paraffin (paraplast+) for 3hrs.

27. Embed tissue in polyfin™ (TBS, # 012 H-PF) paraffin

28. If necessary, soften the faced tissue block by placing in Aerosol OT (Fisher Scientific, # SA292-4) dilute with distilled water 1:10 and immerse for 10 - 20 min.

29. Drain and blot excess Aerosol OT from the block. Cut sections at 5 and float onto a warm water bath.

30. Place sections on an APES coated glass microscope slide.

31. Heat immobilize sections in an 80C oven for 2 hours.

32. Deparaffinze and hydrate to water using standard protocol.

33. Remove carmine stain by immersing the sections in a 0.1% solution of lithium carbonate until the sections are colorless (5-10 min.)

34. Rinse in several changes of distilled water and proceed with routine H&E stain with the following exception: Use Scotts water rather than ammonia water to blue the hematoxylin stain. Ammonia water will cause the tissue to detach from the slide. Dehydrate, clear, mount with permount or other resin