Hi all,

I’ve been trying for the last several months to optimize immunoblotting assays on primary lymphocytes. I am looking at the activation of the NF-kappaB pathway by probing for IkappaB and IKKbeta phosphorylation in activated thymocytes and splenocytes. I have worked out the system with PMA/Ionomycin, but am now looking to use more physiological relavant agonists.

I am stimulating the thymocytes with anti-CD3e and anti-CD28. I have seen people use both plate-bound and in-solution stimulation techniques and I have a range of antibody concentrations (2ug/mL-10ug/mL). Some literature even states 5-10×10^6^ purified cells are used. However, since the phosphorylation events I am looking for are at low stoichiometry, I have to use large numbers of cells (10^7^ per time point). I have tried the plate-bound method with poor results since there is a limit to the number of cells one can stimulate by this approach. I have seen papers use anti-hamster IgG against the anti-CD3 and anti-CD28 to stimulate in solution. I think this is my best option in order to stimulate large numbers of cells.

My question is what is the ratio of antibody to cell number have people used to get optimal stimulation for immunoblot assays? Many references state the total cell number and concentration of antibody, but without the volume in which the stimulation is done, I cannot determine if I am using too little or too much antibody for the number of cells I am stimulating. The same question applies to splenocytes stimulated with anti-IgM?

I know the technique has been used numerous times, but the details are hard to come by in the literature.

Thanks for your input,
-Brock