What is the origin of your antibodies? Mouse or rat or something?

Rabbit...

I incubate the antibodies overnight but trying less time (1-2 hours) at RT the same results...

The blcoking buffers are about standard. Fixation may be an issue, I usually got best results with 4% paraformaldehyde, 10 at RT.

Maybe also the secondary antibodies were too concnetrated. Does diluting them further help with the specificity (without also removing the signal, of course)?

- Fixation with 4% PFA

- Blocking with 5% serum from secondary antibody's host.

- incubation primary antibody 1h RT and then over night 4 degree (antibody concentration !!!!)

- wash 2x

- Incubation secondary antibody 90min RT (antibody concentration !!!!)

When antigens are membrane protein don't use anything which destroys membrane like Triton X-100.

Your antibodies are specific ? and secondary antibody is not crossreactivity?

You may have to use the pre-immune serum from the host animal. A picture of how they look like will be very useful for the discussion.

Make sure your antibodies are rated for IF. There are great deal of antibodies that are rated for paraffin embedded culture or other fluorescence applications but will not produce a satisfactory result for IF on a fluorescence microscope. If you accidentally order a primary meant for a western, for example, your results will be sub-optimal. I have personally never had to use anything other than 1% BSA in 1x PBS. How long are you incubating your secondary? You generally want to also titrate the concentration of the primary and secondary for optimal results - it sounds like you may be using too high of a concentration of secondary. Secondaries are generally not one size fits all. Generally, 1:1000 works fine, but you may want to go even lower.

Hi Alessandra,

To get a good IF staining, proper fixation and permeabilization is very much important along with the other criterion as mentioned above.

What I generally do:

1) wash cells (on cover slips) thrice with PBS.

2) fix cells either with methanol (ice cold, for 10 min) or 3% formaldehyde + 0.025% glutaraldehyde or 4% paraformaldehyde for 20 minutes at RT.

In case of methanol fixation I always get lesser background than the aldehyde fixation. But in some cases methanol fixation destroy the protein structures. So you have to make sure that methanol does not have any effect on your protein of interest.

3) after washing thrice with PBS, permeabilize cells with 0.1% triton X-100 or with 0.05% saponin in 2% BSA containing PBS (that is also used as the blocking buffer) for 30 minutes at RT. For methanol fixation you don't need to use triton X-100 or saponin for permeabilization.

4) after removing blocking buffer, directly add the diluted primary antibody (in blocking buffer) on the top of the cover slips and incubate for 1hr on ice.

5) after washing thrice with PBS add diluted secondary antibody (in blocking buffer) and incubate for 1hr on ice in dark.

6) wash the cells thrice with PBS and then mount on glass slides using mounting medium and keep in dark overnight at RT.

If your antibodies are good then the above protocol would help you to fix your problem.

Best!

Arun

Dear colleges

Well I have the same problem exactly and I need some help!

I am trying to label Bmp2 which bind to the cell receptor in 96 well plate, first I fixed the cell with 4% PFA and second I added BMP2 for two hours at RT, then I added the 1ry Ab ( anti Bmp2 Santa Cruz 1:100 for 1 h at 37 'C for 1 hour then I washed with PBS then I added 2ry Ab but unfortunately I didn't get any signal, I tried 10% Goat serum as well as 1% BSA as blocking buffer but I failed, if any one could help me I would be very thankful for his advice, Best Regards

Did you get any positive result with this antibody in this cell type by any other method, like western Blot?

Ideally, if you have an overexpression construct with a tagged variant of your protein, you can compare the signal from the anti-tag antibody with the anti-BMP2 antibody.

I also generally got better results if I incubate the cells overnight at 4°C with the primary antibody, unless the signal is really cery strong. BSA as a blocking reagents tends to give lower background in IF.

Yes actually I confirmed my antibody with western and I could detect BMP2 , I think that overnight would be good suggestion as last time my incubation was only for one hour .. Thanks for your tip . I will try to use BSA as well in this trial.

There are many good suggestions above. If none of them are not working to you, try with Animal Free Blocker which reduces the background.

When fixation and washing is not proper, backgaround staining may appear.

See: Immunohistochemical localization of mitochondrial fatty acid β-oxidation enzymes in rat testis. Fukasawa M, Atsuzawa K, Mizutani K, Nakazawa A, Usuda N.

J Histochem Cytochem. 2010 Feb;58(2):195-206.

According to my experience the best blocking buffer for the Immunofluorescence is " PBS - BSA 1% For 30 minutes at room temperature, and also you should dilute the primary and secondary antibody with the same blocking buffer

P.S, while preparing the PBS - BSA 1%, you should filter it to remove and contaminant and then store it at 4C

May I ask if I want to monitor both marker located on cellular membrane and cytoplasm, how to optimize permeabilization step so not to damage cellular membrane.

In our lab, some people prepare their blocking buffer with PBS + 5% normal serum (e.g. goat or another - according to your antibodies)+ 0.5% BSA. And that usually works fine. 

I usually prepare my blocking buffer with BSA-PBS 1% and working fine with me (incubation should be 30 to 60 minutes) at room temperature. other wise, I do the dilution for my primary and secondary antibodies with the same blocking buffer.

Hany, could BMP2 be getting washed off? If you aren't adding it until after fixation, then the BMP2 wouldn't be fixed to the receptor, so you could be washing it away. I have never tried labeling a ligand, but you could maybe try adding the BMP2 before or during fixation.

Thanks Dr. Dylan, thanks for all