Hello, I'm having some trouble with my negative control slides when I do immunofluorescence. I only work with frozen muscle tissue, and I'm trying to optimize a number of antibodies. For some of the antibodies, the negative control works ok, but for others, it looks exactly the same as the slides I'm testing! I'm attaching my basic protocol and also some pictures. The first picture is of the MBNL1 protein; this negative has less staining than the test slide. The second picture is of the Palladin protein, and you'll see that the negative is VERY similar to the test slide. For both images, the negatives are on the right side. And for the negative slides, I am doing everything the same as the test slides except I am incubating with PBS only instead of primary antibody and PBS. Any suggestions/tips? Thanks!
You may have an autofluorescence issue to sort out or you may need to titer both your primary and secondary antibodies.
This most likely means that your primary antibody is not working for this tissue. In tissues many antibodies do not work. You may have to do antigen retrieval. You can check the antibody first in a western blot. Then try antigen retrieval.
In my experience I have seen good and bad antibodies. You may also try another antibody.
The signal from primary should be very high if it really works. Don't fall for the false positive results.
Regards
Hi Adrienne, I agree with Peter. To optimize your staining results you have to run a dilution raw. If you have trouble to distinguish autofluorescence from the real staining result you can change the color, instead of CY2 or alexa488 use CY3 or alexa 546. I recommand if you are not working with double labelling you also can detect your anitbody binding with DAB. You can use either Peroxidase labelled secondary abs or the Avidin-BiotinComplex System or Streptavidin in combination with biotinylated secondary abs. Good luck!
More information is needed to figure out the problem.
1- Did you use same acquisition settings for both images?
2- Are you using same secondary antibody for all stainings? (maybe using goat serum for blocking is not the right choice for all your secondary antibodies)
3-Is your Palladin antibody tested for IF-frozen tissue and PFA fixation?
I recommend you to optimize primary antibody stainings using well known positive control tissue or cell lines. Adding 0.05% tween20 to the washing buffer (especially during and after secondary antibody incubation) might be helpful. Fixation is also an important factor some antibodies work better with acetone fixation.
Bleach and check again. If it stays there it is autofluorescence.
Also the glycogen deposits in the muscle fibers are meticulously autoflorescent.
You may also try to detect your epitopes using red and far red fluorescence where the auto is diminished.
Cheers,
Ivan
I titrated out my secondary antibody and I am still getting lots of staining with the negative control slides. I'm using a goat anti-mouse Alexa and blocking with 5% goat serum. It's extra weird to me because I've done this same stain before and didn't have this bright staining with the negative controls. I'll check for fixative-induced fluorescence next, but if I didn't get it before, why would I get it now? It's the same tissue and same protocol. So frustrating!
What species does your tissue come from? If it's mouse, then you may have issues with non-specific binding of your secondary antibody. You can try the mouse-on-mouse (MOM) kit to help, or switch the species of your antibodies.
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