BTW, half log units would be 1:10, 30, 100, 300, 1000, 3000, etc etc. With the procedures outlined in the paper attached to the previous note, I have found NO antibodies which are used at concentrations of more than 1:1000 so that is where I start. Moreover some antibodies I use have optimums at 1:3,000,000 (yes million)! Two were monoclonals, one a polyclonal so one must carry the range far enough to see it decline. With fluorescence, using direct tagged secondaries would have failed at 20-50x higher concentrations. A few of the weaker monoclonal antibodies would not have worked at all with direct tagged fluorophores but did work well for ABC immunoperoxidase. That said use of short incubation times with primary antibodies can require 10-100x more antibody, use of TSA amplification for fluorescence enables use of primary antibodies easily 50x-100x more dilute - their optimum is frequently = to or just slightly higher than that we obtain using ABC immunoperoxidase with NiDAB chromogen.

Hi Bodhisattwa,

I usually prepare all antibodies with different concentration from 1:100 to 1:3200 in 1% normal goat serum in PBS to find the best concentration. I think it is the best way to save your time and find the best concentration.

Good Luck

Hi, unfortunately the titration of the antibodies depends on the conditions of your experiments, I think that you have to try different dilution.

I can recommend you that most of the primary antibodies works at the concentration of 100ug/ml. But probably you have to adjust the concentration according to your experimental condition.

Good luck

Barbara

Hi,

First of all I check whether there are some suggestions in the datasheet ( this helps to lower the amount of testing:)0

If there is nothing mentioned, I normally start with only 3(-5) dilutions:

1:100

1:500

1:1000

(1:2000, 1:50)

After testing this, I can narrow down to an optimal concentration.

The secondary antibody should be always the same for better comparaison.

Good luck!

Hi,

I agree with the previous answers. I also start with few dilutions in PBS: 1:50, 1:100, 1:200,1:500, 1:1000. This gives you a good range to work with and optmize secundary antibody concentration. But everything depends on your application.

Good luck!

Hi,

Different dilutions in light microscopic ICC may help you to choose an optimal dilution for fluorescence ICC.

All the Best!

Concentration of antibody depends on tissue. The best way is testing range of 1/100 to 1/ 2000

Good Luck

hello, As suggested above setting up dilutions over a range, say 1:10 to 1:2000 would help you narrow down. Also you can include a blocking step after the primary ab incubation. Wash adequately for neat background. Good luck.

Thanks to all of you.

These are all good answers. One other thing to add it that it can help to have good positive and negative control cells/tissue so that you can be sure what is real labelling versus any non-specific signal

Run a positive control (Which should be an antibody which you know your secondary fluorescent antibody will bind too) and run a negative control (Which should be a non specific antibody) and run your sample. Put up around 5 dilutions covering a wide range like 1:10, 1:100, 1:1000, 1:10000 and 1:100000. Do this for all of the above mentioned samples. You should get no florescence in your negative control. This will help you decide a narrow range of dilutions for your next experiment. Best of Luck!

I agree somewhat with Kripa's evaluation but find it more useful to use increments of half log units rather than full log scales. Moreover one should NOT in my view titrate an antibody using fluorescence but rather should obtain the optimal concentration with ABC immunoperoxidase methods first, then convert that number to the correct concentration for fluorescence. use of secondaries directly tagged to a fluorophore is not only the LEAST sensitive of methods, but is more difficult to tell if your range is perfect or not. Please see the attached article on titrating antibodies

BTW, half log units would be 1:10, 30, 100, 300, 1000, 3000, etc etc. With the procedures outlined in the paper attached to the previous note, I have found NO antibodies which are used at concentrations of more than 1:1000 so that is where I start. Moreover some antibodies I use have optimums at 1:3,000,000 (yes million)! Two were monoclonals, one a polyclonal so one must carry the range far enough to see it decline. With fluorescence, using direct tagged secondaries would have failed at 20-50x higher concentrations. A few of the weaker monoclonal antibodies would not have worked at all with direct tagged fluorophores but did work well for ABC immunoperoxidase. That said use of short incubation times with primary antibodies can require 10-100x more antibody, use of TSA amplification for fluorescence enables use of primary antibodies easily 50x-100x more dilute - their optimum is frequently = to or just slightly higher than that we obtain using ABC immunoperoxidase with NiDAB chromogen.

Strange qu !

One of the responders mentioned data sheets. companies do not tell you which method they used or the conditions for staining. In my experience the "data sheet" is used only as a measure of the highest concentration you would use in titrating. Thus if the company mentions 1:10,000, then I start there and go to 1;10,000 1:30,000 1:100,000 1:300,000 and 1:1,000,000. Always include 3 log scales. a simple check if your method happens to be better than what the company tested over just a 10 fold range will be essentially useless in my experience. The half-log point is 3x more dilute and thus I like using that number as well as the intermediate value. I have also learned NEVER to use short incubation times for the detection step or the time of primary incubation. Counter to logic, primary antibodies do best with more than a 24 hr incubation with the tissue in the cold or require very very high concentrations to get any binding. This is an issue even with tissue mounted on slides. The incubation with the fluorescent tagged secondary I find works best at 37˚ for 2-3 hrs as well. Conc. usually 1:600-1:1000.

I always look at the data sheet to find a starting concentration. Then if the company recommend 1:50, my first run will have 3 concentration 1:25, 1:50 and 1:100. The reason why I go higher than the suggested concentration is because I want to make sure I go signal. At the beginning it does not matter to me if it is perfect. For titration purposes I always do overnight 4 degrees. Another very important factor at least to me is Epitope retrieval. It matters a lot! If you archived a good epitope retrieval your titration becomes easier. No all antibodies will give you a great signal with 6.0 pH retrieval. I hope you can archive your goal with the answers and different points of view. Let us know how was it. Good Luck!!

Thank you everyone for your answers. I have tried with a dilution of 1:500, 1:1000, 1:1500 and 1:2000 for the primary antibodies and 1:500 of the secondary cy3 conjugate. But unfortunately I am still finding quite a good amount of background. So I guess I'll try with more dilutions. Please let me know if I should dilute my secondary antibody any further?

Which antibody are you testing?

Hi,

probabely you have to change your blocking conditions?You should always block with the serum in which the secondary was raised, then block in a 5% solution for 1h and I even dilute in these cases my 1.ab in this blocking solution.

Next, I think 1: 500 for the secondary antibody is quite high, I normally take 1:1000 or even 1:2000.

Have you tested the secondary ab in another IF?

It is also important to have a negative control ( sec. only) to see unspecific binding.

Good luck!

Correct, having a positive tissue is of benefit. When examining whether a new molecule is present is a bit more challenging and knowing if there is biochemical evidence of the molecule in the tissue is then more critical.

Sorry to say, I don't think this question can be answered in a forum such as this. There are many variables to consider and details about antibodies, target antigen, tissues, are missing. For optimization the correct positive and negative controls are essential, for both Abs and tissues, and a criss-cross titration may be necessary. Choice of fixation, blocking reagents, diluents, incubation time and temperature, etc, may all have an impact. There are plenty of methodology papers and manuals to consult - some of the better ones are published by companies which sell the antibodies. To get optimal results, it may be necessary to do a few preliminary experiments using controls, varying this and that, to home in on the final conditions.

I agree with Gerald. The matter is too complex to be completely solved here. Bodhisattwa what kind of tissue are you staining? What antigen is your target? W"hat fixation and what antigen retrieval method you are using? Without these info all answer are superficial and unsufficient. Anyway, it not important the starting dilution reported on the data sheet, but the Ig concentration of the antibody. I always look at the data sheet to find it and then I start with 2-5-10-20 ug/ml Ab concentration in PBS (or TBS) containing the blocking medium (BSA 1% or Normal serum in which the second Ab is made). Similarly, the second antibody was used at 2 or 5 ug/ml for 1 hour at RT.

Repeated washings are needed before sealing sections.

If you have too high background,possible causes can be many. here it is hard to imagine them all.Let we know.

One thing to keep in mind is that titration of an antibody with direct fluorophore secondary antibodies is much more difficult than using chromogen-based staining and converting the results to fluorescence. See the attached document which points out WHY. Moreover the concentration needed for the primary antibody is dependent upon the time of incubation with the priimary. The shorter the time, the higher the required primary. I agree with a previous comment that the secondary concentration we use is a bit high but "if it ain't broke, don't fix it. We have used 1:1200 with success so it is probably not a critical feature. Time with primary is, however important. We are now comparing different times (with freely floating material mainly but have used paraffin section as well and >24 hrs gives more detection even in relatively thin paraffin sections. Access is the problem for sections on slides whether frozen on paraffin and we find that longer rinsing times may be needed for mounted sections as well. That does not mean that everything has this problem as better antibodies = more forgiving conditions.

Oops here is the paper