What is the best container in which to put a tissue sample to fix in 10% Neutral buffered formaldehyde? Can I use just a centrifuge tube or should I use a glass bottle?
A glass bottle may brake. It is better to use a wide mouth plastic container with hermetic seal
You can use both. Be sure to close any one of them tightly, and if you are going to store tissue for long time, you have to change the 10% Neutral buffered formaldehyde at regular base.
I suggest plastic container with wide cap and at least 10-20 ml capacity, because of slow perfusion of formaline. in my own experience in glass container the part of tissue sample in contact with glass doesn't fixed probably result of adhesion! so its better to use plastic one
Thank you. Dave, I will ask my lab staffs to be aware of your suggestions.
Dear Sulie, As i am used to work with the histological sections. Most of time i put 10% neutral buffer formaline in glass bottles / plastic bottles. I never felt or know any problem with these both materials.
If you know anything or get anything good please do share that detail.
Regards,
Dear Mansur,
The glass bottle was what I was instructed to use many years ago. I also interpretated that there must be some reasons why glass bottle was used. However, I do not know if there are problems in using plastic bottles since I have never used them for formaline.
Regards,
I personally think ,the volume of the fixative is more important. it should be 15 to 20 times that of volume of specimen.Glass/ plastic bottle doesn't affect much....preferable glass bottle but now a days plastics are more commonly used.
As my colleagues suggested before I agree with them and my self I used both plastic and glass bottles and no problem has happened. The good thing that the spcemin quite small I mean not very thick so formol can penetrate and then good fixation of tissues and the volume about 50 times the volume of the specimen and we have to chake the bottles sometimes after fixation and at the coming hours in order to homgenize the the fixation at least 2-5 times a day at the first few days. If you want to kkep the samples for a long time I think you save them in 50 or 70% alcohol (Ethanol). good luck.
Any container made of inert material is OK. Any plastic is OK, and is not essential. The other things are way more important. Ensure the formalin is 10% neutral buffered, and the volume tissue/fixatiove is at least 1:10, time is also important, average optimal is 24h; any underfixation is usually a bad thing- therefore nobody recommends to fix for less than 8 h even small tissue fragments. Some structures are sensitive to overfixation, so more than 72 h is generally not recommended.
I think an inert plastic container sould be the most appropriate for delicate studies, as immunohistochemistry. It presents two advantages over glass containers: 1) it doesn't break under casual shocks; 2) certain types of glass have ions in its composition and those ions could dissolve in the fixative solution, altering its properties. However, in routine histology/histopathology, both containers (plastic and glass) are suitable, though plastic containers are physically more stable.
I really appreciate your thoughtful insights about the ions of some glass composition. Thank you.
I use 10% phosphate buffered 10% formalin for my processing, and I leave it in for at lest a week. I've found the buffered version to be less toxic and gentler on the tissue.
If you are using plastic tubes or flasks without a gasket I recommend you to test that they are not leaking before using them with formalin (leave a few containers upside down filled with water for a some hours). I ve had quite a few disappointing experiences with "high quality" products and you want the formalin to stay inside the container. We re now using small bottles of the HDPE-type and they are good but expensive...
Tubes are usually not recommended as the tissue could end up in the tip and this may cause lack of fixation.
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