I have been doing immunostaining of cultured cells. The current issue is that I don't see any positive stain. One thing that worries me is the fixation and permeabilization step. Is it possible that the protein (both in cytoplasm and nucleus) got lost or washed away during the steps? I used ice cold acetone for 10 min RT. I have been searching the fixation and permeabilization steps online. The are pretty similar. Has anybody ever had this experience before? The same procedure works fine in tissue sample. It's tricky for cultured cells on slide.
Check the antibody spec sheet for fixation tips. Some antibodies work well with formaldehyde/paraformaldehyde fixation, but not methanol/acetone, and vice versa. Most likely due the binding domain recognised by the antibody, it can be altered during fixation. Whether the protein is extracellular, or intracellular of course also influences the staining protocol you use. Your problem may not be of fixation, but of inadequate permeabilisation, but then again, it seems you got it working on tissue sections, which is usually the harder of the two. Have you tried a range of fixatives and triton concentrations in e.g. a multiwell chamberslide, so that you can optimise the staining?
Are you using an established lab protocol, or are you troubleshooting a protocol? If it is the former, look back to your lab notebook and try to identify where you could have erred. If you are troubleshooting (which seems to be the case), the process can be painstaking. You do not want to shock your cells and degrade your epitope. You may have to change the incubation times, or your dilutions. Tweak these factors in very systematic ways and see what works. I wish you the best of luck. I am attaching a very good IHC paper which may or may not be of use.
Hi Zhenning, I agree with Michael in that antibody reactivity can differ dramatically depending on how you fix cells.
Acetone and organic solvent fixation techniques usually strip away the lipid membranes from cells. If your antigen(s) are small proteins, then it is likely that they can also get washed away.
Try doing fixation using 1% PFA for 20 min in the presence of 0.1% Tween-20 or TX-100 without agitation of your slides and see if that helps.
You don't mention which antibody(ies) you are using, and whether or not it has been shown to work with acetone-fixed tissues. Is this the first time you have tried the ICC on your cultured cells?
If you are using acetone fixation it is unlikely you would need to perform further permeabilization as the acetone will, as Steingrimur says, remove the lipid component of the cell membrane.
You also have to be wary of acetone in that it can introduce shrinkage artifacts in cells and tissues (it is a potent dehydrant), and does not preserve tissue and cellular architecture well. Acetone in primarily employed when surface cell markers are used (e.g. lymphocytic markers)
Previous experiments I have conducted with cultured skeletal muscle and neurons show that IF usually works well with a minimal fixation in 1% paraformaldehyde of 5 minutes. In addition the briefer the fixation time, the less likelihood of having to perform any antigen retrieval.
However without knowing the specifics of the antibody you are using I cannot offer any more advice
Thank you all for the suggestions. I have been using a rabbit primary antibody from Dako. This is my first time staining cultured cells myself. I will watch the fixation time carefully. Another thing I have seen is that many fixing steps are done at -20C with ice methanol, or even acetone. Is it necessary to fix cells at -20C with formaldehyde? As you have pointed out, the antigen could be distorted after fixation if not washed away. I will try to do a gentle fixation and permeabilization. I read that 0.5% Tween20 is more gentle compared to 0.1% Triton /NP40. Will I be able to stain nuclear proteins with Tween20 permeabilization? Thanks!
Hi Zhenning, it could be that you are not using the best primary antibody for your studies.
Dako has great products for labeled secondary's, lectins, etc, but they are not well known for their primary antibodies.
I suggest that you look for another primary antibody that recognizes your protein when fixed and what fixation procedure works. Here are some antibody search engines:
http://www.linscottsdirectory.com/
http://www.antibodydirectory.com/
http://www.antibodyresource.com/findantibody.html
hi, i am also making a cell culture for mast cells, usually it is in falsk. first I put cells on the slide using cytospen on polylysin slides.
for fixation i used 2 fixatives so you can try them as well.
first and best: modified Bouin’s fluid [95% saturated picric acid, 5%
concentrated (37%) formaldehyde, and 2.5% glacial acetic acid] for
10 min at RT, permeabilized in absolute methanol for 10 min at RT,
and stored in 70% ethanol at 4°C until further processing for immunocytochemistry.
Second use PFH4% for 10 min followed by aceton/methanol 50/50 for 10 min,then leave to dry in air for 15 min and store in -80.
this is for the fixation problem I hope this helps you.
In vitro activation of murine DRG neurons by CGRP-mediated mucosal mast
cell degranulation
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