I am working on setting up a fragment screen. I am able to successfully immobilize the protein on various chips (NTA capture, NTA couple+capture, SA chip) at various densities (1000-8000 RUs). The protein is active in solution: binding of various small molecules by DSF and ITC give thermal shifts/KDs in the expected range. However, once immobilized onto the chip, I see at best 6-8% surface activity which is insufficient to run a fragment screen.
Has anyone else come across a situation where the protein is active in solution but not when immobilized on the chip? Any suggestions for how to get this assay to work?
Here are a few things I have tried:
1. different constructs:
shorter and longer versions of the same protein
3. different temperature
25C, 37C, 15C
4. different buffers
This is really puzzling since the tags should not interfere with the protein activity. Here some thoughts:
Biotinylation can take place at the binding site inhibiting subsequent analyte interaction but that is not expected with the HIS-tag.
Sometimes there is a negative correlation between amount of immobilized ligand and ligand activity (Zhao, Gorshkova, Fu and Schuck; A comparison of binding surfaces for SPR biosensing using an antibody–antigen system and affinity distribution analysis; Journal/Methods; (59) 328-335; 2013) meaning that there is an optimum in the ligand immobilization.
Maybe you can immobilize the protein while already interacting with an analyte. In this way the binding site is 'protected' during the immobilization.
Could it be that the dextran is interacting with your protein and thus 'inactivating' it?
I found one publication about heparin immobilization in various ways resulting in different surface activities. (Osmond, Kett, Skett and Coombe; Protein-heparin interactions measured by BIAcore 2000 are affected by the method of heparin immobilization; Journal/Analytical Biochemistry; (310) 199-207; 2002).