Hello there,
I am having mixed results with golgi-cox staining and I was wondering if anyone has experience/insight into the individual stepwise process. My issue is that the neurons are not looking as they should.
Ideally, I would like to avoid buying the rapid golgi-stain kit. I have little interest in spending a month processing tissue so the ultra-rapid golgi method is preferred.
Thus far my protocol has been developed from the following two methods papers:
https://www.ncbi.nlm.nih.gov/pubmed/29155038
https://www.frontiersin.org/articles/10.3389/fnana.2019.00062/full
That said, the tissue I am working with so far has been 1X heparinized PBS perfused mouse brain (looked correct under the microscope) and PBS/4% PFA perfused rat brain (I wind up with bushes/exploded looking things.)
My impregnation solution is 1% each concentration of mercury chloride, potassium chromate and potassium dichromate where the whole-brain (rat) is submerged in the impregnation solution for 36 hours at 42C (PFA-perfused).
Tissue is then put in cryoprotection solution (25% sucrose/15% glycerol) for a minimum of 24 hours.
Tissue is then frozen/embedded in ice using a mold in our cryostat set to CT -24C/OT -23C to create smooth slices (tissue was previously shredding when sliced despite using a new blade in the cryostat.)
Tissue is then mounted to a ice basement on the puck.
Tissue is cut at 150um thickness.
Collected slices are placed on 2.5% gelatin coated slides and adhered with 30% sucrose. Excess sucrose solution is absorbed off the slides/tissue using filter paper and gentle pressure. Slides are then left overnight at 30C in a slide warmer or placed in our incubator set at 37C for ~10 minutes if I am feeling particularly lazy.
Development is as follows: 20 minutes in 30% ammonium hydroxide followed by 5 minutes in distilled water followed by 20 minutes in 10% sodium thiosulfate (I am aware that these concentrations are high but I have yet to read why certain concentrations are chosen in various method papers in order to properly adjust for my protocol).
Our dehydration sequence is 5 minutes each of 50% ethanol, 70% ethanol, 95% ethanol, and 100% ethanol. Lastly, xylene is used twice for 5 minutes each wash.
Tissues is covered with DePex mounting media, coverslipped, and sealed - stored away from light until viewing. Slides are typically viewed the day after.
Additionally in some tissue (mouse), we attempted PACT instead of CLARITY and attempted to enhance the GFP-tagged cells with IHC (this was just for kicks to see if we could, because if you can do CLARITY with golgi and you can do IHC with golgi, why not try all 3 at once...)
**The problem is that instead of getting happy little trees with long branches, I am getting fluffy bushes. Fluffy, exploded, sad bushes. Things that one cannot analyze because they are trash.** See attached photo - apologies for having to take it via cellphone through the objective. Couldn't get a detailed image through the scope's camera.
Does anyone have an insight onto what is causing this issue/what adjustments to make to my protocol to avoid this problem?
Hi! I think that: 1. the impregnation time is too short even if the results are important, necessary, urgent or curiosity is so big; 14 days in Golgi-Cox solution I think is necessary. 2. NH3 and thiosulfate are too concentrated; if you use 14 days for impregnation, 3:1 NH3/Water, and 5% thiosulfate you obtain good results. Anyway, my protocol based on the few articles is: 14 days impregnation, paraffin embedding, sections 25-50 um, hydration, NH3 conc. 8 min, water 2x5 min, thiosulfate 5% (avoid light) 10 min, water 1 min, ethanol 96%, 5 min, 100% x 2, 5 min, xylene, 5 min.
Hi,
I face the same problem, I followed the protocol in the following paper:
Article Golgi-Cox Staining Step by Step
However, I perfused the rats with Accustain to replace PFA. Do you think accustain affects the impregnation? accustain is alcohol based fixative!
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