Hello everybody,
I have been working for several years on sea urchin gonad development using histological techniques, now I am going to start working on the histology of fish larvae (Pagrus pagrus and Mugil cephalus).
I only have this detailed protocol, which is perfect for sea urchin gonad:
Fix in 10% buffered formalin
Wash with tap water
Store in 70% ethanol
1h 70% ethanol
1h 85% ethanol
1h 95% ethanol
1h 100% ethanol
1h 100% ethanol
1h 100% ethanol : Bioclear (1:1)
30' Bioclear (room temperature)
30' Bioclear (oven, 60-62°C)
Paraffin overnight (60-62°)
3h Paraffin (60-62°)
Embedding in paraffin, room temperature
Cut with Leica microtome, oven 39°C overnight
20' bioclear
20' bioclear
20' bioclear
3' 95% ethanol
3' 90% ethanol
3' 70% ethanol
3' 50% ethanol
5' distilled water
30' haematoxylin
quick wash in distilled water
20' tap water
5' distilled water
3' eosin
quick wash in distilled water
quick wash in 90% ethanol
3' 95% ethanol
5' 100% ethanol
5' 100% ethanol
20' bioclear
20' bioclear
20' bioclear
Closure with Eukitt
In you opinion, can it be good for fish larvae (from 1 to 50 dph)? How can I improve it? I would like to specify I don't have an automatic tissue processor, so I usually do all the passages manually. I read a lot of protocols using tissue processors but they are too long, therefore they cannot be used in a manual procedure.
Can you help me? Thanks in advance,
Barbara
hi Barbara
as you know larval histology is really hard and you should do it careful.
you said that
1-Paraffin overnight (60-62°)
in my experience on Zebrafish and sterlet sturgeon larvae: in num 1 decrease temperature to 56 (it is good for paraffin melting: higher temperature will change shape of your larvae!)
i will send you some useful link, also if you want aid you can say me personally.
A high-throughput histoarray for quantitative molecular profiling of multiple, uniformly oriented medaka (Oryzias latipes) embryos
High-throughput zebrafish histology
kind regard
Buffered formalin is a good general preservative but not the best histological fixative. You might consider switching to more standard histological fixatives that better "fix" cell and tissue structure and are more adaptable to specific histological stains. Often mercuric fixatives give good results but there are many to choose from depending upon your specific needs. For general staining procedures (such as haemotoxylin and eosin) Zenkers fluid works well as does Holland-Bouins. I suggest you check a standard histotechnique methodology source for approaches that best adapt to your ultimate microscopic needs.
Stardard dehydration, paraffin embedding, sectioning, and staining with H&E should be okay for larger fish larvae as larvae will lie flat in a square mold form in molten paraffin. But for smaller and especially yolk-sac larvae, I recommend using plastic, ie glycol methacrylate, to embed specimens. A standard pharmasutical capsule can be used as a mold. A good staining combination for plastic sections is alkali blue 6B - neutral red (H & E will not penetrate the plastic).
Dear John, unfortunately I do not have the possibility to use methacrylate as embedding agent. I have only paraffin available. So I need to adjust my usual method for larvae histology.
Unfortunately you don't say what you specifically want to study, and depending on what you want to do, different techniques may be appropriate. As mentioned above, the common method of embedding larvae these days is to use the water soluble resin glycol methacrylate which is polymerised using UV light- but it was not always so. Formalin/ paraffin has the disadvantage that the tissues shrink and distort more than in the methacrylate. Low temperature paraffins are available, or freeze drying, or even resin embedding for TEM, cut at 3 microns with a glass knife will work, but finding a suitable stain is an issue. I have usually just used formalin/paraffin and H&E stain, but then I am usually just looking for lesions, not producing an atlas or a development paper. My suggestion is that, if paraffin is the only option available that you look at the old literature from before 1970. Use Google Scholar with the custom range set to exclude any modern papers and see what the early researchers used. You will find they often used fixatives such as Zenkers or Bouins as mentioned by Dr Prezant above but increasingly histologists are reluctant to use the hazardous chemicals involved, even though they are superior to formalin (bouins in my opinion is unsurpassed for gonads). As an example, early methods for very small larvae included using vital stains, clearing the larvae and mounting them whole (no sectioning).
Dear John, I would like to study the development of the larvae. I will look for old literature. The early method you spoke about in your last sentence is very interesting...
The biggest issue for me in cutting the very large salmon larvae was the yolk sac. 8 micron sections and high humidity (breathing on the sections). Super sharp blades are a must. I used buffered formalin and the specimens were processed with a machine that put them into wax (no xylene - an orange based solvent). To me its all about the cutting - having a good set up and the tissue in a solid block for slicing. If it was dry, the yolk shattered.
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