ER stained with Ab looks bad... - (Mar/30/2009 )

I have tried to use an antibody from AbCam (ab13504) that recognizes calnexin in ER. My cells are HEK293, and I've fixed them in either cold (4oC) methanol for 10 minutes or in 2 % paraformaldehyde for 15 min at room temp., then incubated with PBS with 20% FCS, 5 % BSA and 0.1 % saponin at room temp. for 30 min for blocking/permeabilisation. Both primary and secondary antibody was diluted in this buffer (incubated about 50 min at room temp with both antibodies with several washes in PBS in between). But the resulting ER staining is very dotted, and extremely different from what I see when I transfect these cells with a plasmid expressing an ER-localized protein with fluorescent tag (this is in live, unfixed cells). I didn't expect the antibody to be as pretty as the "ER-transfected" cells, but these dots don't look very good. They are mainly located in the cytoplasm and usually not so much in the nucleus (which is good), and they are not due to background staining from the secondary antibody.

Is there anybody out there that has tested this antibody and got nice-looking staining? Or perhaps someone has suggestions about how to improve my fixation and/or immunostaining protocol? Thanks in advance

Sonjaa,

While I've never used this antibody, I do have a suggestion... Did you try preabsorbing your antibody? While you may be sure that the background isn't from your secondary, it could be nonspecific staining by the primary itself. Preabsorbing your antibody will eliminate much of the nonspecific binding, and usually is more effective at eliminating the nonspecific binding than just using FCS in the antibody solution.


-Carlton

Carlton H on Apr 5 2009, 05:28 PM said:

Thank you for your suggestion, Carlton. I have done some more testing now, and it seems that these membrane structures are very sensitive to fixation. The appearance of the antibody staining (as well as the appearance of the GFP-tagged protein we are studying and which we believe is retained in the ER) is very much affected by the fixation protocol. Reducing the temperature of the PFA solution combined with the use of saponin to open the cells makes the staining look even worse. The problem now is not dots but rather vesiculation. So, cold PFA followed by saponin gets really bad (by the way, cold PFA alone without saponin makes our GFP-tagged protein look beautiful, just as it appears in live cells, but is obviously useless for antibody staining since the cells are not opened). Cold MeOH is better, but then the staining of the antibody is very "dotty". So, my theory is that at lower temperatures, the PFA fixation is not complete and the membrane structures get destroyed when we add saponin. Also, someone in this forum has mentioned that cells might be able to do a lot of different things during the first minutes after adding PFA and especially if the solution is relatively cold (this will stress the cells and the fixation goes relatively slowly). I will try PFA at different temperatures to see how this affects the cells. Also, some others here in our lab is using PFA first for just 5 min then followed by cold MeOH at -20oC for 20 min and no saponin or any other detergent (not necessary, since the MeOH will open the cells). I'll try that, too. If that doesn't help, I may have to try to improve the antibody as you suggest. But until then I will stick to the idea that I have a fixation-problem and not an antibody-problem. We'll see...
Thanks again!

Another fix to think about is 3% TCA (trichloroacetic acid). Not commonly used, but we've had success using this when PFA fixing failed (in IHC applications).

Good luck,
-C