FW: [Histonet] Re: Pam Marcum colleague losing bone sections from slides

Tue Apr 7 17:37:48 CDT 2015

-----Original Message-----
From: Carol Fields 
Sent: Tuesday, April 07, 2015 3:37 PM
To: 'Cooper, Brian'
Cc: 'histonet-bounces <@t> lists.utsouthwestern.edu'
Subject: RE: [Histonet] Re: Pam Marcum colleague losing bone sections from slides

I've used that also many times.  It really does work.
Carole Fields,  HT (ASCP)
Lead Histotechnologist, Pathology Laboratory Martin Luther King Jr. Community Hospital
 1680 E. 120th Street
Los Angeles, CA 90059

-----Original Message-----
From: histonet-bounces <@t> lists.utsouthwestern.edu [mailto:histonet-bounces <@t> lists.utsouthwestern.edu] On Behalf Of Cooper, Brian
Sent: Tuesday, April 07, 2015 3:23 PM
To: gayle.callis <@t> bresnan.net; histonet <@t> lists.utsouthwestern.edu
Subject: RE: [Histonet] Re: Pam Marcum colleague losing bone sections from slides

Have you ever tried blotting your slides dry before putting them into the oven?  The veteran who taught me this trick used it on brain tissues from the Coroner's office (in the early 80s--don't wanna offend anyone) which were grossed very thickly and were always poorly processed.   She never used charged slides or additives in her waterbath.  She claimed she was the only one in her lab who was "allowed" to cut this stuff because everyone else's slides had tissue loss!  I can tell you from my experience that it works well for toenail which is notorious for detaching from slides.  I've used it on many other tissues as well.    

Anyway, press a slightly moistened clean L'Absorb or paper towel down onto the section after microtomy.  You don't want the paper towel soaking wet--just damp.  This will effectively wick the section of any excess moisture.  Then incubate and stain as usual.          

Good luck.

Brian D. Cooper, HT (ASCP)CM | Histology Supervisor Department of Pathology and Laboratory Medicine Children's Hospital Los Angeles
4650 Sunset Blvd MS#43- Los Angeles, CA 90027 bcooper <@t> chla.usc.edu 

-----Original Message-----
From: histonet-bounces <@t> lists.utsouthwestern.edu [mailto:histonet-bounces <@t> lists.utsouthwestern.edu] On Behalf Of Gayle Callis
Sent: Tuesday, April 07, 2015 2:57 PM
To: histonet <@t> lists.utsouthwestern.edu
Subject: [Histonet] Re: Pam Marcum colleague losing bone sections from slides

>From Pam:  I am currently trying to stain L6 vertebrae from rabbits. 
>They
have been decalcified and paraffin processed properly. I've tried cutting at both 5 and 10 microns and my tissue is still not sticking to my slides. I know my sectioning is fine because I'm successful with every other tissue I've ever sectioned and stained. For some reason the bone I'm using won't stick to any slides. I was using charged slides and I even tried poly-L-lysine slides, but the bone keeps coming up even before I attempted to stain them. I've even tried leaving them in the incubator for more than the usual 48-72 hours. I know it's possible to do other stains beside H&E on bone, but I think my main issue is just getting good contact between the tissue and slide. If you have any advice or thoughts, I would love to hear them. 

I will get the messages to him ASAP. 

Pam 

*************************************************************************

What was meant by incubator and at what temperature?   It helps to dry
sections FLAT, at 37 to 40C for several days.  Do NOT dry at 60C.   

If the sections are not staying on plus charge or poly L lysine coated slides,  then use chrome gelatin subbing solution in a water bath OR by
pre-subbing clean microscope slides.    

This is the Chrome gelatin protocol that worked for our huge decalcified bone sections and or problem bone sections. 

Chrome Gelatin Subbing Solution:  Section/Slide Coating Adhesive

0.1 g Chromium Potassium Sulfate (this is toxic.  Collect for proper disposal, not down the drain is you pre-sub the slides). 

1.0 g Gelatin:  100 bloom, Sigma.  For large bone sections, use 200 or 300
bloom gelatin, Sigma).   200 and 300 bloom gelatins are very large gelatin
molecules made from pig collagen.  100 bloom is a much smaller molecule than
200 bloom.   Do         NOT use household (cooking)  gelatin used for
cooking. Buy the pure gelatins only. 

1 liter Distilled Water

                Dissolve chromium potassium sulfate and gelatin in hot but not boiling water.  Cool subbing solution before use, and store in refrigerator.  If gelatin gets growth, discard, make new.  A few crystals of Thymol in stock subbing solution can help prevent growth. 

DO NOT USE PLUS CHARGE SLIDES WITH SUBBING SOLUTION.   GELATIN COATS OVER A
PLUS CHARGE COATING AND NEGATES THE POSITIVE CHARGE.  

For presubbing glass slides, wash these by dipping in acetone, air dry before using the pre-subbing protocol to get rid of any greasy/oily residues
on glass surface.   If you put the subbing solution in a water bath,
uncoated,  glass slides will work fine without further washing.    

You can do either of the following: 

1.            Add 10 ml subbing solution to a warm water bath for paraffin
sections. Then mount sections onto the cleaned glass slide, drain, and air dry, store in a cool, dry place. 

2.            Dip acetone washed, dry slides into subbing solution, air dry,
and store in a dust free area.  Box subbed slides and store until needed.  

If you get background staining with hematoxylin (hematoxylin stains gelatin) then dip  pre-subbed slides in NBF ~10 times, rinse with distilled water, air dry and store slides.  The aldehyde fixative cross links the gelatin to some degree, but still allows section to adhere without annoying background staining.  

Pick up sections from water bath drain and lay flat to dry at 40C for
several days.   You will not need extra subbing solution in the water bath
if using presubbed slides. 

IF all else fails, try Sterchi tape transfer method with packaging tape.   I
have the method with photos and publication, and will send privately.   

Good luck

Gayle Callis

HTL/HT/MT(ASCP)

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