Tissue is fixed in 10% NBF before being vacuum embedded in paraffin. Our tissue
is cut at 10microns, placed on charged slides, then placed on a slide warmer
over night. The slides are then place in xylenes 3 times for 2 minutes, then
stained as follows..Step 4: 100% Alcohol – 2 X 2 minutes each,Step 5: 95%
Alcohol – 2 X 2 minutes each,Step 6: DI H2O – 2 X 2 minutes each,Step 7:
Harris Hematoxylin – 1 X 1.5 minutes,Step 8: Wash gently in DI H2O until“Grape
Juice” color is gone, Step 9: Acid Alcohol – 3 Dips, Step 10: Wash gently in
DI H2O – 1 X 2 minutes, Step 11: Bluing – 10 Dips, Step 12: Rinse in running
DI H2O – 1 X 2minutes, Step 13: 95% Alcohol – 1 X 2 minutes ,Step 14: Working
Eosin – 1 X 2 minutes, Step 15: 95% Alcohol – 2 X 2 minutes each, Step 16:
100% Alcohol – 3 X 2 minutes each,Step 17: Xylene Dips – 3 X 5 minutes each,
Step 18: Coverslip.
Our core lab has recently had a change in pressure for the DI port and water
comes out very hard, making gentle washing impossible. The reagents are new.
We have tried increasing the staining time in the hematoxylin to 2 minutes and
reducing the acid alcohol dips to 2.
Our hematoxylin is not consistently staining the nuclei in the ligament tissue.
Some are good, some are bad.
Can someone make any suggestions?
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