I am currently working on binding affinity assays using MST for homologs of the plant LecRK receptor for NADP (LecRK6.2-2). Previously, binding was assayed using P32 labeled NADP, and was found to have a Kd of ~1 uM.
Now, when trying to assay binding using MST, we are running into some issues. The protein is expressed and purified in E. coli using a C-term His-tag purification, and is fused N-term with Sf GFP. We were originally measuring GFP fluorescence, but acidity from NADP was interfering with the signal. Now, we are dying the His tag with RED-tris-NTA for the reporter. When the protein level (ligand) is varied against constant dye (target), we get a beautiful curve. When the dye level is varied against constant protein, it flat lines.
The question is, is the small (nearly insignificant relative to protein MW) change in MW after dye binding not detectable by the MST machine? If so, is there a way to optimize for small ligand binding, as we intend to test NAD(P) binding?
It would be helpful to attach images of your capillary scan and thermophoresis curves. I'm assuming the flat lining isn't the instrument having its sensors saturated?
Without seeing this data, I'd say if your his-tag dye shows nanomolar affinity to your target and you're running the protein in something like 2x molar excess, you should move on to optimizing your assay by: analyzing the thermophoretic trace of labeled protein alone (or with positive control single concentration ~10xKd) for replication of curves, buffer optimization and MST setting screening if the first experiment has unacceptable output.
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