Immunocytochemistry - staining for cytokeratin - (Sep/10/2008 )

I am currently staining fixed luteal cells for cytokeratin using a fluorescently tagged primary antibody. Other people in the lab have used this antibody before and we are sure it works. I decided to stain slides that had been fixed for a couple of months and stored in the fridge and I got nothing! I kept trying and trying and nothing seemed to work. So, the lab got fresh tissue, dissociated it, and started fresh cell cultures. I fixed a slide and immediately stained it for CK and it worked. However, we tried to stain another slide 2 weeks later and it didn't work. We are using 2% paraformaldehyde and post-fixing with ethanol. The antibody is only about a month old, aliquoted and stored at
-80. My advisor is confused because he stained fixed cells months after the fixation and the staining worked - the images are even on the poster outside our lab! Is there some reason why over time the fixation would deteriorate and the antibody would no longer work? Has anyone had similar problems?

Is your paraformaldehyde (PFA) freshly made up? PFA goes off very quickly, I usually have single use aliquots frozen in freezer or in the fridge for a week at most. Also check the PFA is correctly made up too. Heating to 70C PBS etc. etc. Your original fixation may affect staining.
Storing cells is ok usually for a short while, but I find after 2 months the results are not so reliable. How do you store them? is it different from your supervisor?
I usually store cells in PBS with BSA and Sodium Azide sealed with parafilm in the fridge. If the samples have been fixed with PFA I store them after fixing but before extraction with alcohol.
For cytokeratin staining I usually fix cells with 1:1 Methanol: acetone ice cold solution for 10mins instead of using PFA (no need for post processing either) as I find that gives much better defined filaments under the microscope.

Is your paraformaldehyde (PFA) freshly made up? PFA goes off very quickly, I usually have single use aliquots frozen in freezer or in the fridge for a week at most. Also check the PFA is correctly made up too. Heating to 70C PBS etc. etc. Your original fixation may affect staining.
Storing cells is ok usually for a short while, but I find after 2 months the results are not so reliable. How do you store them? is it different from your supervisor?
I usually store cells in PBS with BSA and Sodium Azide sealed with parafilm in the fridge. If the samples have been fixed with PFA I store them after fixing but before extraction with alcohol.
For cytokeratin staining I usually fix cells with 1:1 Methanol: acetone ice cold solution for 10mins instead of using PFA (no need for post processing either) as I find that gives much better defined filaments under the microscope.

I have been making the fixative fresh, letting the slides air dry, then placing them in the fridge with parafilm wrap. I have tried storing them in PBS and another wash buffer that has sodium azide, and that seemed to help the staining a week post fix (I stained a slide that was fixed and stored our normal way and there was no staining). My advisor was concerned that storing the slides in a buffer would deteriorate the fixative -- is this a valid concern? We are getting cells today, so I am planning on testing out these other methods!

No I think the storing will not affect the fixative, Curtis is right usually PFA is used at 4%, 2% seems quite low.
Are you cells grown on the slides? I try and not let the cells or tissue dry out (unless is just after sectioning frozen sections)as that affects the staining (I get alot more artifacts if they dry out).

Its always good to try comparisons, I hope it works for you.

Let us know

Lost

I will be growing some HeLa cells next week, so hopefully I'll have some answers soon...I'll keep you all posted. thank you SOOO much!