[Question from NMRWiki Q&A forum] Observing protein-ligand interaction using weakly soluble compounds

[nmrlearner]

Observing protein-ligand interaction using weakly soluble compounds

I am trying to setup an experiment using HSQC with titration of a compound to determine the Kd of the protein-ligand interaction. My difficulties have been in the sample preparation/formulation, as these novel compounds are very weakly soluble in aqueous buffers. I must be able to quantify the soluble concentration of compound over ~6-8 points to generate the binding curve (signal intensity/chemical shift vs. [L]).

Previous observation of this protein by HSQC and other NMR techniques have used a buffer containing 5 mM HEPES, 1 mM DTT, 5 mM EDTA in 90% H2O/10% D2O, pH 7.5. Compounds have not been soluble in this buffer, and only slightly more so in this buffer + 5% DMSO.

In each of the following methods of solubilization, compounds were successfully dissolved in an organic solvent, but delivery into an aqueous buffer suitable for the protein have yielded opaque solutions with precipitate material.

(a) Dissolve dry compound in DMSO, then add small volume of compound stock to NMR buffer.

(b) Dissolve dry compound in CHCl3, deposit on glass vial surface, then add buffer and sonicate and/or place in ultrasonifer water bath.

(c) Dissolve dry compound in concentrated HCl, then add buffer drop-wise.

I would appreciate any answer to the question of how to prepare a sample using weakly soluble compounds such that I can calculate the Kd of the protein-ligand binding.



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