I have one issue in my antibody internalization experiment. I added antibodies to incubate with cells on cell culture plate for 2-6 hr ,and then fixed/permeabilized cells and used secondary antibodies conjugated Alexa 488 to detect the cellular location of incubated antibodies. However, I found there is very strong fluorescence background on my plate. It seems the incubated antibodies bind on the bottom of cell culture plate.
Have anyone had the same problem? Please let me know if you have any suggestion to improve.
Thank you
You can try to increase the stringency of the washing steps (augmenting time, or tween concentration, or even changing the washing buffer)
Also try to reduce the concentration of the secondary antibody, or the incubation period, or the temperature of incubation.
Does your protocol include some blocking step? Check it carefully. Augment the time or concentration of your blocking agent.
Hi, Rosa:
Thank you for your useful suggestion. I will add some tween in my blocking and wash buffer. So far, we only use PBS to wash.
Thanks again.
You can try to use higher cell density (close to or more than 100% confluence) before the adding your primary Ab to the cells. In that case you will decrease the surface of interaction between the primary Ab and the unoccupied surface of the cell culture plate.
Yu-Chih, do you include a blocking step (e.g. 1% BSA or 5% NFDM in PBST ) before you add your secondary antibodies?
Actually, I'd expect the surface of the dishes to be blocked already by the serum proteins during cell culture, but exactly there could be the root of your problem: Cross-reactivity of bound serum IgG with your detection system, easily to be confirmed by background in not antibody-treated controls. If that should be the case, you could try IgG -depleted serum (e.g. home-made by passing serum through a protein A/G column) or establish a detection system that does not interact with (I'm assuming) bovine IgG.
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