FACS antibody titration - (Jan/10/2012 )

Hi there,

I will be titrating my FACS antibodies (and this is my first time). Can anyone please answer some very basic questions below?

- Which cells should (or can) I use to titrate? (I will be sorting primary cells from normal tissue and/or normal epithelial cell line. Can I use ANY cells?)
- Is there any case where Fc-blocker is not required?
- Does the procedure somewhat vary depending on conjugation status (flourochrome, biotin, or non-conjugated), and if so, why?
- Do you recommend serial dilution or adding different amount of antibody to the same volume of buffer?

Thanks!

sc_queen on Tue Jan 10 06:35:21 2012 said:

1. You can use any cell you want to do a titration but you have to ask yourself what are you optimising for. If you are detecting say, marker A on a specific cell line B, then the wise thing would be to use cell line B + antibody against marker A. However, this is only the case if you know cell line B expressed marker A (this can be done by literature search, gene expression experiments, immunostaining etc.). The flow antibody spec sheet usually comes with a 'positive control cell line' which will definitely express your marker of interest (it is usually immortalised cell lines or isolated mammalian cells).

2. Fc blocker is usually used on mouse/human serum analysis where you test for immune cell subpopulations. The rationale is that secondary antibodies could potentially bind to primary cells hence giving a false positive background. The main thing is to have a matched isotype control (this is the Ig of your primary antibody - again can be found in the spec sheet). It is advisable to use the same concentration as your primary and then use this as the 'background reference'.

3. If you're using the primary antibody for the first time, I would recommend you titrate the primary and secondary. Although most of the time, small changes to the secondary dilution doesn't affect the final data much (but who knows, don't risk it).

4. This is totally up to you. I normally just add the appropriate volume in the flow tube. E.g. 0.1, 0.5, 1.0, 2.0, 4.0 ul into 100ul cell suspension. I find this easier to handle if you have alot of tubes.

My 2 cents...

You should use cells (or a cell line) where only a subset of cells express your protein. This way you can distinguish positive events from negatives.

Fc blocker is required for Fc receptor positive cells, such as macrophages, lymphocytes, monocytes, so all lymphatic preparations and blood cells. If you there are no such cells in your tissue, you don't need to use Fc blocker.

Antibody titration should be done for any antibody, independent of conjugation.

Thanks Science noob and Rsm! It really helps.

How do you analyze your data to determine the optimum concentration, by the way? If you could point me to a good website, that would be helpful, as well. My search has not been very successful... Thanks!

sc_queen on Mon Jan 23 16:31:07 2012 said:

I find it easiest to understand if you ask yourselfwhy are you even doing titrations in the first place. A few questions and answers I posted to myself which may help you answer that question:

1. To make sure your antibodies do not create a false positive - that is to say, negative cells should not appear stained due to overwhelming amounts of antibodies

2. To make sure your positives do not become a false negative - that is to say, all positive cells are all fully stained

3. To use less antibodies - if all your positive cells are fully stained, and increasing the concentration of the antibodies does not improve the percentage, you can save some antibodies

4. Good separation is a bonus - a positive peak, and a negative peak

There is actually a graph to guide you to find your optimal concentrations.. but I can't find it now. Vaguely, it looks like an X, and the optimal is somewhere near the middle of the X. The axis are probably X=antibody concentration, Y=signal intensity. With that in mind, hopefully you can determine your optimal concentrations.