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Fixation of HEp-2 in immunofluorescence slides - (Oct/13/2010 )

I'm doing a protocol for detection of ANA using HEp-2 cells and immunofluorescence. But I've got a problem! The slides, after fixed, has to be kept in the freezer, and I need them to stay in the refrigerator.

Anyone knows of any preservative or fixative which allows me this?

I use the following protocol:

Grow 3 ml of Hep-2 cells in 12 ml of DMEM. Incubated in stove of CO2 at 37 degrees.
I released the cells with trypsin and trasfer to other two bottles.

Then, I apply 50 ul of the HEp-2 cell in the slides,and Incubus in stove of CO2 at 37 degrees by 37 to 24 hours. Remove of the stove and fixed with methanol and acetone 1:1.


I'm confused as to why you say the slides have to be kept in the freezer. The methanol/acetone fixation requires you fix the cells at -20 degree for only 10 minutes. After that you can take them out and keep the in the refrigerator. Is there something special about your cells that I'm not aware of?? Anyway, the other option you have is paraformaldehyde which fixes at room temp but it doesn't maintain specific cellular structures such as centrosomes.


Thank you very much for your answer!

My problem is that after fixation with methanol/acetone at -20 degrees, the cells in the slides degrade when stored in the refrigerator, keeping only its stability in the freezer. But I need it to remain stable in the refrigerator.

How do that other treatment with paraformaldehyde you mentioned?


Oh...I've had this problem. Are you just using uncoated glass? I found that when I cultured my cells on collagen-coated coverglass they remained on the glass much better. How quickly are your slides degrading? As for the paraformaldehyde, you can buy the powder and make 3.7% paraformaldehyde in PBS but this is time consuming and you have to heat the mixture up for it to go into solution. Rather, I buy 16% paraformaldehyde solution (EMS Cat#15710) that I dilute with PBS to 3.7%. You just put the paraformaldehyde on your cells for 10-20 minutes at room temp and they are fixed. You then need to permeabilize your cells by treating with 0.5% TX-100/PBS for 10 minutes before blocking. Here is a link that gives you the protocol to many different fixation methods: Perhaps one of these will suit your needs better.


Yes, I'm using uncoated glass, maybe here is the problem! You buy the collagen-coated coverglass ready to use or you make it in lab? If you make in house, how do you do that? My slides are degrading about 1 week!
So, I'll try to use the collagen-coated coverglass and if it don't work I'll try to change the fixation.

Thank you very much for your help!

I await your return!




We buy our coverglass already coated with collagen. Check out BD Biosciences. They offer coverglass coated with many different proteins and you may find that collagen is not your best condition. They offer poly-D lysine, poly-L lysine, laminin, fibronectin and collagen. If you call them, they may be willing to send you out a couple samples so you can test which will work best for this specific cell line. You can try to coat your own coverglass, there are many protocols online, but I've found this to be difficult and the results are highly variable. Nunc also makes coated cell culture chambers. I've found the cells adhere much stronger to these but the problem is the slide is plastic and large. You get higher background because of the plastic and it takes much more antibody to stain the entire slide. Otherwise, you might need to process your samples faster and not let them sit in the refrigerator for a whole week. For one experiment in particular, I found that my cells were no good after about a week (even with the coated glass) so I had to stain and mount them as fast as possible. I found that after staining and mounting I could keep them at 4 degree for weeks without any problems. That way the sample was good and I could analyze the staining at my convenience.