I have come across a few protocols using different increments of ethanol, but most of these refer to tissue sections. Most others in our lab have also done this with tissue sections. I am using a cell line, and I really want to avoid disturbing the proteins in or around the plasma membrane if at all possible.
I use to use 3 types of fixation method for IF:
1) 4% PFA in PBS
2) 3-4% Formaldehyde + 0.025 Glutaraldehyde in PBS
3) ~100% ice-cold Methanol
In case of PFA or Formaldehyde/Glutaraldehyde fixation, I generally get higher background than that of Methanol fixation. To get less background I use to use 0.3M Glycine in blocking buffer (2% BSA containing PBS). For this you have to permeabilize your cells. I generally use 0.05% saponin or 0.1% Triton x-100 in blocking buffer.
Some antibody staining works well in PFA or Formaldehyde/Glutaraldehyde than methanol fixation and vice versa. The type of fixative to use and the condition of fixation mostly depends on antigen being studied and the antibody being used to stain.
Fixation with methanol denatures proteins, so never use if you have a fluorescent protein that you want to look at in your sample. Some antigenic epitopes are in fact completely destroyed by methanol. However, it might also be advantageous for some other antibodies (especially some monoclonal antibodies which bind only one epitope) that the naturally buried antigens could be exposed after methanol fixation.
For IF staining of lipid bodies one could not be able to use methanol/acetone as fixative.
So, what kind of fixative should you use mostly depends on molecules (antigens) of your interest being studied.
Hope this could help you.!!
Best!
Arun
I use to use 3 types of fixation method for IF:
1) 4% PFA in PBS
2) 3-4% Formaldehyde + 0.025 Glutaraldehyde in PBS
3) ~100% ice-cold Methanol
In case of PFA or Formaldehyde/Glutaraldehyde fixation, I generally get higher background than that of Methanol fixation. To get less background I use to use 0.3M Glycine in blocking buffer (2% BSA containing PBS). For this you have to permeabilize your cells. I generally use 0.05% saponin or 0.1% Triton x-100 in blocking buffer.
Some antibody staining works well in PFA or Formaldehyde/Glutaraldehyde than methanol fixation and vice versa. The type of fixative to use and the condition of fixation mostly depends on antigen being studied and the antibody being used to stain.
Fixation with methanol denatures proteins, so never use if you have a fluorescent protein that you want to look at in your sample. Some antigenic epitopes are in fact completely destroyed by methanol. However, it might also be advantageous for some other antibodies (especially some monoclonal antibodies which bind only one epitope) that the naturally buried antigens could be exposed after methanol fixation.
For IF staining of lipid bodies one could not be able to use methanol/acetone as fixative.
So, what kind of fixative should you use mostly depends on molecules (antigens) of your interest being studied.
Hope this could help you.!!
Best!
Arun
If you fixed properly your cells considering the membrane proteins, dehydration after staining should not cause problem.
Then: after staining your samples, dehydrate with increasing ethanol series. defat in xylol. embed the coverslips by DPX, for example (or any other anhydrous embedding medium). The protocol which i use is the same, but shorter in time, then others use for tissue samples (due to the monolayer of cells, they thin). Usually i dehydrate the coverslips in wells, for xylol, i use a short deeping by holding one coverslip with forceps. thats all.
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