ICC: High fluorescence cell body staining - secondary staining all neuron cell bodies (Sep/28/2005 )

Hi,
I'm having trouble with my immunofluorescence of primary cultured neurons.
My secondaries, both FITC and Texas Red, brightly stain the cell body, regardless of the type of cell. I generally culture the neurons on coverslips, fix in 4% PFA for 15 min, and permeabilize with .25% or .1% Triton-X for 5 min. I've tried changing the conditions of primary and secondary antibody incubation with little change.
However, when I use a tyramide/rhodamine amplification kit (that detects a HRP conjugated secondary) I don't have this problem at all!
Does anyone know why the fluorescent secondaries stain the cell body and how I might stop it.
Thank you!

What are the details of your protocol so we can help ... like what do you block with? Are you sure your antibodies are specific and titrated properly? Are you doing double staining? If so are your single stains ok?

I've tried a couple different protocols, all similar except the triton-x.

4% PFA fix for 15 min RT
followed by block with 5% NGS in .1% Triton-X for 1 hour at 37C
then a primary o/n incubation at 4C (primary is in PBS only)
next day 3washes 10 min each in PBS
secondary incubation 1 hour at RT
then 4 washes 10 min each in PBS
mount with vectashield

another protocol is:
4% PFA fix for 15 min RT
permeabilize in .25% Triton-x in pbs for 5 min RT
block with 10% NGS in PBS for 1 hour at 37C
then primary o/n incubation at 4C (primary in 3% NGS in pbs)
next day 3washes 10 min each in PBS
secondary incubation 1 hour at RT
then 4 washes 10 min each in PBS
mount with vectashield

I've tried altering the above protocol by adding 0.05% Triton-x to the washes-with no difference.

I'm not sure how to check the titration, we had the primaries made, the secondaries are from Sigma and Chemicon.

It's single staining of a sodium bicarbonate transporter, NBC. We have 2 primary antibodies for the transporter, one in rabbit and one in guinea pig. I worked with our rabbit NBC in the past, using a goat anti-rabbit FITC secodary. Now I'm trying out our guinea pig NBC with a goat anti-guinea pig Texas Red secondary. WIth both sets of staining I get strong cell body staining that stains all neurons, almost like the bodies are glowing.
However, when I use an HRP/tyramide/rhodamine amplification kit with the rabbit primary, (that antibody is very weak and limited supply) I get beautiful specific staining only along the membrane and neurites.
I don't get it.

This tells me that PROBABLY your secondaries are the problem as the tyramide kit uses a different secondary than the homemade protocol right? What concentration do you use the secondaries at? Do you run controls with isotypes or no primary? You probably do not need to do overnight incubation of primary also. And make sure to dilute your primary in block or a diluted form of it. This is important!! Also are the secondaries very specific? I get my secondaries from Jackson ImmunoResearch as they are the best! I use 1ug/ml for 30 minutes diluted into my block or a diluted form of it. I also use donkey instead of goat as I get more background with goat.

This is a tricky one!

Yeah, this is science torture.

I have a correction to make on the above protocol, the anti-guinea pig Texas Red is made in donkey (I used NGS incorrectly)

Ok,
Yes, with the tyramide kit I use a different secondary, goat anti-guinea pig HRP from Chemicon. Using this kit with this secondary, I get background staining when I omit the primary. Because of the background staining, I moved to a fluorescent conjugated secondary IgG, Texas Red, from Jackson Immuno, concentration of 1:200 or 7.5ug/ml.
I have tried 1 hr room temp primary incubation, but my signal was super weak. The primary is not the finest quality, not purified and weak.
I have heard conflicting opinions on diluting my primary in a diluted block versus PBS alone. I've tried both, (but I may have used the wrong species in my block as in the above protocol). I will repeat Monday using the correct species as a block.
I'm not sure about the specificity, my secondaries are just IgG, no subtypes.

Thank you for your help!

Being that the antibody was made in donkey and you used goat serum shouldn't be a problem as long as your antibody is cross adsorbed. Otherwise it is a good idea to switch serums. Look up the catalog # on Jackson's website and see if it's cross adsorbed.

The background staining you initially got with the HRP secondary probably is from endogenous peroxidases, were these blocked when you were doing that protocol?

You only need to use the secondaries at 1-2 ug/ml at most, believe me. Above 3-5 ug/ml is too too much. These are polyclonals and must be diluted out.

I would definitely try a series of dilutions/concentrations for your primary.

What do other people tell you about diluting the primary in block or some form thereof? To me it is a basic thought process in all immuno experiments as you want to be sure your primary will not non-specifically bind to any agents in your block.

i get the same thing with my cytocpins, block with BSA, use a rabbit produced antibody, a FITC conjugated secondary and voila... my negative controls are bright green!

If you want to provide more details of your experiment I would be glad to try to troubleshoot with you as well. But with the info you have given it's too difficult to think out the multitude of possibilities.