Dear Sera
 
Sounds like you need more advice with muscle stains. Ph is extremely
important for the ATPase.  Your protocol doesn't include the preincubation
solution either. The Gomori's stain is easily wiped out with too strong a
differientiation solution. We don't even use one it takes away the red stain
that is so important to see the mitchondria with. The PH of the Gomori's
solution is also very important it is different than the regular paraffin
Gomori Stain.
 
How are you freezing these muscles?
 
I think you need much more information than I can go into on this email but
if you really are going to proceed and be successful in this endeavor I
would be willing to send you a muscle handout that was written by Peggy
Wenk. It is our bible of sorts. Let me know.
 
Annette Featherstone HT/MLT
 
 

-----Original Message-----
From: Sera116@aol.com [mailto:Sera116@aol.com]
Sent: Monday, April 21, 2003 19:11
To: histonet@pathology.swmed.edu
Subject: Frozen Muscle STains


Hi Everyone, Thanks for the responses, I appreciate it. Heres my questions: 

how long does a frozen muscle have to be in the -70 freezer before you can
cut it on the cryostat? 

Do any of you use control slides when staining? And if you do use controls
do they have to be fresh tissue or paraffin embedded tissue? 

The stain giving us the most problems is the ATP. We can't seem to get it to
work consistently. We also are not doing the pH, and wonder how important
that is to the staining process? I'm thinking pretty important. 

The staining protocol we are using is as follows: 
1. Slides in Sodium barbital buffer for 10 minutes. 
2. incubate slides in incubation solution at 37 degrees c. for 30 minutes. 
3. Transfer slides to 2% cobaltous chloride solution for 3 minutes. 
4. Wash slides in distilled water for 5 minutes. 
5. we then dip slides (one by one) in Ammonium sulfide 6-7 times, sometimes
more. I believe the stain gets darker the more you dip. 
6.Rinse in water and check differentiation. At this Point I expect to see
two different shades of brown. 
7. Wash in running tap water for 20 minutes, if everything looks like it
should. 
8.Deydrate in graded alcohols, clear in histoclear or xylene and coverslip
in synthetic mounting medium. 

The solutions are made the following way: 

Sodium barbital buffer: 
sodium barbital 2.062g 
calcium chloride .999g 
d. water 1000 ml 
The ph is then supposed to be adjusted to 9.4. Stable for 2 months. Store in
refrigerator. 
We do not do anything with the pH because this is not how we were taught to
do the stain. 

2% Cobaltous chloride: 
Cobaltous chloride 2.0g 
d. water  100 ml 

Ammonium sulfide: 
Ammonium sulfide 1.0 ml 
tap water 10 ml  (but I was taught to use 20 ml) I don't know why. Does this
need to be made fresh each time? Should the Ammonium sulfide be stored in
the dark? 

Incubation Solution: 
Sodium barbital buffer 10.0 ml 
Adenosine-5-triphosphate (ATP) 15mg 
This also needs a pH of 9.4 and is prepared fresh each time. Again no pH
modifications are done. 

Basically when I do this stain it never looks the way it looks in the
textbook. It always looks mottled. It never looks crisp with a distinct
color difference. 

Does this stain react differently with different muscles? If the muscle
sample is of poor quality does it affect the staining? 

I am also having trouble with the Modified Gomori Trichrome: 

how important is it to stain the slides within one hour of cutting? 

does anyone filter the trichrome solution before staining? 

I find this stain very basic yet it never comes out consistently. As of
late, there is a dark , black  precipitate like artifact on the slides after
staining. I'm not sure whats causing it. 

Heres the protocol: 

Trichrome Solution 
Fast Green FCF  0.03g 
Chromotrope 2R 0.6g 
Phosphotungstic Acid 0.6g 
glacial acetic acid 1 ml 
d. water 100 ml 
Mix well and adjust pH to 3.4 ( again no pH adjusting here) 

Stain slides in Harris Hematoxylin for 5 minutes. 
Rinse in Distilled water  to remove excess Hemo. 
Stain in trichrome solution for 20 minutes. 
wash in distilled water to remove excess trichrome. 
Dip twice in 0.2% acetic acid. ( we sometimes use 3%). 
Dehydrate in graded alcohols and clear in xylene. Mount in synthetic medium.


I'll get back to everyone on the Nadh, Pas, Pasd, oro later. I don't want to
bombard everyone. Feel free to post these on the histonet if its easier for
you. Thanks to everyone who responded. I appreciate it.