I am working on determining the complex structure of a membrane protein with a drug compound.
In attempting to obtain the complex structure, I believe there are two main approaches: imaging the sample in a micellar state or reconstituting it into a lipid membrane, such as a nanodisc. How are these methods typically chosen or differentiated in practice?
Are there any cases where a binding site was not observed in the micellar state but became visible after reconstitution into a lipid membrane?
IMO you should start by reconstituting in detergent because it’s the fastest and easiest way, with the lowest sample loss. Speed is often critical with less stable proteins. A detergent suitable for high efficiency extraction can be used during lysis (along with gentle mechanical disruption), and then replaced during affinity purification or SEC by different detergents more suitable for structure determination or more stabilizing to the particular target. Generally speaking ultra-low CMC detergents such as LMNG and GDN are best for structure determination, while moderately low CMC detergents like DDM are best for extraction.
Nanodisc reconstitution can provide a more native environment (though not necessarily as the lateral membrane pressure is higher than in a regular membrane). Some proteins are significantly more stable or more active in nanodiscs, and indeed there are a number of cases of lipid-protein interactions, re-entrant domains, or lipophilic ligand sites that are better or only resolved in nanodiscs. The cost is additional prep time, and typically loss of ~50-70% of the material during reconstitution.
Other options are amphipathic polymers like PMAL-C8 and A8-35, native nanodiscs with SMA or AASTY polymers (SMALPS), Saposin A lipid nanoparticles (SaLiPro), peptide-based soluble adapters (Peptidiscs), traditional proteoliposomes made by extrusion, and native membrane vesicles produced directly from cell membranes and concentrated by ultracentrifugation.