Author Topic: Issue with cresyl violet staining of rat brain sections  (Read 6082 times)

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Offline PDavid

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Issue with cresyl violet staining of rat brain sections
« on: January 27, 2016, 01:46:33 PM »
Hey all,

We have been having some issues with sectioning and staining rat brain tissue.  We are doing doing a simple nissl + myelin stain.  Here are all the details:

(1) Rats are perfused with 4% paraformaldehyde.  After perfusion, brains are post-fixed in PFA for at least 24 hours, but some of the brains have been sitting in PFA for up to 1 year.  Brains are then transferred to a 30% sucrose solution, and allowed to sit in sucrose for at least 72 hours (although some brains have been sitting in sucrose for up to 1 year).

(2) Brains are removed from the sucrose solution, and the first 10 mm of brain tissue is "blocked".  The whole brain is embedded in TissueTEK embedding medium (we also have Shandon M-1 available, but have typically used TissueTEK).  The brain is sectioned in a cryostat set to -21 C.  Tissue sections are 40 um thick.  We save every tissue section from the 10 mm block of tissue.

(3) For mounting tissue, we have tried 2 different techniques.  The first method is to take each tissue section and store it in a small well of 0.01 M PBS until we are ready to mount the tissue on slides (typically less than a week, often only 1 or 2 days).  The other technique is to "direct mount" tissue in the cryostat by lowering a slide down to the stage and "catching" the tissue on to the slide.  Often times, when we use this second techique, we wet the slide with a little bit of PBS because I have found it allows the tissue to "catch" easier, and as far as I can tell it helps eliminate bubbles in the tissue.  I will refer to these two techniques as "wet mounting" and "direct mounting".  "Wet mounted" slides are mounted using a dish of PBS and a paintbrush.

(4) Tissue sections are mounted on SuperFrost Plus slides.  No gelatin needed.

(5) After mounting, slides are typically allowed to air-dry at room temperature for a day or two before staining.  Occasionally staining happens on the same day as mounting.  Also, occasionally, staining happens up to a week after mounting.

(6) The staining protocol is thus:

2 min dH2O - hydrate/rinse tissue
2 min 70% EtOH - dehydrate tissue to prepare for clearing
2 min 95% EtOH - same as above
2 min 100% EtOH - same as above
5 min Citrisolv - clear the tissue (we use Citrisolv instead of Xylene)
2 min 100% EtOH - rehydrate tissue to prepare for staining
2 min 95% EtOH - same as above
2 min 70% EtOH - same as above
2 min dH2O - same as above
10 min Myelin stain (we use a Cyanine R-based myelin stain, pH is 2.0)
1.5 min differentiation step (in an ammonium hydroxide solution)
2 min dH2O
20 min Cresyl Violet stain (0.1% cresyl violet acetate solution, pH is 4.5)
2 min dH2O
5 seconds 70% EtOH (dehydrate tissue for clearing)
5 seconds 95% EtOH (same as above)
5 seconds 100% EtOH (same as above)
3 min Citrisolv (clear tissue in preparation for coverslipping)

(7) Coverslipping - We coverslip using DPX as our coverslipping medium.  We do not allow the slides to dry before coverslipping.  We remove them directly from the Citrisolv and coverslip immediately.

So, now that I have described our methods thoroughly, here is the problem: We are getting a lot of horizontal cracking (primarily during the sectioning process), and a lot of curling and sections falling off of slides (during the staining process).  We have tried quite a bit to alleviate the problem, but not much seems to work. 

It seems like the tissue sections falling off during the staining process may have been due to the direct mounting technique - or so we think. The explanation we have come up with is that the embedding medium was getting inbetween the slide and the tissue, and then causing the tissue to fall off during the staining process.  Is this logical?  We have stopped direct mounting, and that problem seems to have stopped - hopefully.

We still have a lot of curling of tissue during staining, and cracking of tissue during sectioning, however.  Cracking usually happens near the caudal/rear portion of the 10 mm block of brain (when we approach the hippocampus), but rarely towards the anterior portion of the brain (motor cortex).  Lately we have attempted sectioning at warmer temperatures (-16 C) as well as tried embedding only the base of the brain instead of the whole brain.  Results are mixed so far.  Also, nothing seems to have helped the curling issue during staining.

I have uploaded sample brain sections to here:

If that link doesn't work, try this one on Google Photos:

Thanks for any advice and comments.  We are really stumped as to how to solve the problem!

Issue with cresyl violet staining of rat brain sections
« on: January 27, 2016, 01:46:33 PM »

Offline OBarbara

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Re: Issue with cresyl violet staining of rat brain sections
« Reply #1 on: May 15, 2018, 01:47:07 PM »
Did you ever resolve the "curling" issue in your sections? I am experiencing the same curling with most of my sections, which are mouse brains, perfused with 4% para, cut on the vibratome at 70um, mounted on gelatin-subbed slides, air dried overnight and then run for Nissl. The protocol I use is the same as yours except the initial time in H20 is 10min, the time in Citrisolv is also 10min, and  the time for Nissyl stain is up to 1min. We then rinse twice in dH20, followed by 2min in 70% ETOH, differentiate for 4min, then 2min each of 95% ETOH and 100% ETOH, then twice for 5min in Citrisolv. I'm wondering if I need to shorten my times to mirror your times, but if you are still having curling issues, then I'm not sure that would help.

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