I have done paraffin-embedded rat brain sections before however in order for all my tissue to be processed the same and for all my antibodies to work, I cannot use paraffin-embedded brain tissue.
My main issue: the brain stains so heavily and no amount of alchaol or lithium oxide will differentiate or remove it from the grey matter. I don't know what to do. De-fat the brain? (Any ideas how to defeat successfully?)
Does anyone have a working protocol?
All my tissue is processed in this way (no way to change it now, what's done is done):
1. overdose of pentobarbitol
2. saline transcardial perfusion
3. cut the rat brain in half (anterior and posterior, with a coronal slice to divide the two) and put into 50mL falcon tubes filled with 40mL of 4% PFA and stored overnight/24hrs at 4 degrees
4. washed a little with PBS and put into 30% sucrose till dropped to bottom at 4 degree
5. put into fresh 30% sucrose in 4 degree till I need to freeze
Now the more flexible part: as I am troubleshooting all antibodies and stains before going ahead with my cohort of brains.
Freezing:
flash-frozen in dry ice cold isopentane in OCT within a mold and then put into the -80 freezer
Slicing: cryostat 30um thick free-floating sections.
LFB staining attempts:
trial 1: mounted the slides and left to dry overnight, washed in dh20, no xylene or Ethanol steps and put into Luxol fast Blue stain overnight in 60degree oven. Next day: 10 dunks in 100% ethanol, then lithium carbonate (ended up leaving in for entire weekend)l tap water and I didn't progress further because the whole brain was dark blue and no amount of lithium oxide or alcohol would remove the stain from the grey matter
trial 2: learnt that for non-paraffinised tissue i should 'de-fat' the tissue. So I did 1 of two things. 1 slide went into a 1:1 solution of chloroform:100% ethanol for either 4 hours or overnight.
for the 4hrs in de-fat solution: Then rinse off in 100% ethanol (5mins), then put into 3 steps of xylenen (5 min each), then rehydrate 100%->70% ehtanol (2 min each). then put into LFB for 1-2hr, then 10 dips in 100% ethanol and water (2min), then 70 to 100% ethanol, and it seemed to take all the stain off (cannot remember my notes aren't that good)
for the overnight de-fat solution: the same steps as above except the stain came off in the first dips after the LFB stain (10dips) and the lithium oxide made no difference to the stain, and the outcome was that 4/5th of the tissue was stained heavily dark blue (no stain was removed from the grey matter) apart from 1/5th of it where the differentiation worked well. I have no idea why..
Does anyone have a working protocol for fixed frozen rat brain tissue?
I have tried IHCworld's and it doesn't work and I emailed them and they just told me to use paraffin....
I've not done this particular stain myself, but something I would think about trying if Luxol blue is essential to your analysis is to use a very dilute Luxol blue solution and stain for a shorter period of time. You might try cutting the Luxol Blue concentration to about 1 tenth of the concentration used in the standard protocol. Trying a slightly lower incubation temperature (instead of 60 degrees C, try 40 or 45 degrees and do a staining time course and pull out samples every 30 or 60 min to check for the intensity of staining until you find the "sweet spot" This is complete guesswork on my part.
Alternatively, could you achieve the result you need for your experiment using cresyl violet, toluidine blue, or some other stain for your purposes or is use of Luxol Blue critical to your analysis?
My lab has done some immunolabeling combined with Toluidine blue counterstaining in the past. If the Tol blue is too intense, you can place the sections in warm PBS for a while to get some de-staining. depending on what IHC markers you are trying to use you may find that the histological stains could mask some of your immunolabeling, so some care and balancing of IHC and histological stain will be important. In our experiments, we've done the Toluidine blue labeling first followed by fluorescence IHC)
Finally, are you using a fluorescent immunolabeling approach or a brightfield sort of approach with an opaque reaction product (like DAB)?
I hope there is something useful for you in this. Good luck.
My expierence is that the LB (Kluever-Barrera-Staining is not suitable fot thick frozen sections. If you need it, I would recommend as it was already mentioned, reduce the concentration of LB, the incubation time and the temperature. And do it step by step. A better counterstain for IHC is Haematoxylene (DAB) or Nuclear Fast red (DAB-Ni-Cb). For IF DAPI.If you need the information of myelinated nerve fiberes I would recommand a double labeling with Myelin Basic Protein (MBP)
Hi All,
Thanks for your time and input. I spoke to a technician in my lab and she found this paper (one i had found but not read yet, annoyingly): https://link.springer.com/content/pdf/10.1007/s00418-001-0357-z.pdf
The method I did which was incredibly successful was this:
Aim: is to test the de-fat step to see if these fixed frozen free-floated then mounted slides will de-lipid enough to have good LFB staining
Slices are mounted free floating slices 30um thick and left to dry overnight on superfrost slides.
Day 1:
· 70% ethanol overnight room temp (20hrs)
Day 2:
1. Then LFB for 6hrs? (not more than 16hr) in oven – 5 works
2. dip into 100% ethanol
3. dip into 100% ethanol (two different clean containers)
4. Then wash in dh20 to get the ethanol off
5. Then lithium carbonate should be seconds differentiate (10-20 dips)
6. Dh20 wash for 10mins
7. Check microscope and see if you need to differentiate more
8. Then put into counterstain (33sec in hameotoxylin)
9. Running water
10. Check stain on microscope
11. Then 90% ethanol dip
12. 100% 2 min
13. 100% 2 min
14. Xylene 3*5mins
15. coverslip
it worked very well on my test naive slide
Hazel G May Thanks for posting the protocol that you found was working. I am just testing different conditions since it's our first time to do this in frozen sections. I have one final question, in the final version you mention that the aim is to test the de-fat step, how did you do it? alcohol chloroform 1:1? or just directly 70% ethanol, dry, LFB stain?
Thank you
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