Biological organisms rely on proteins to perform the majority of the chemical tasks needed to sustain life. All functions that proteins perform stem from the three-dimensional structures that they adopt. Because these structures are stabilized by weak non-covalent interactions, at temperatures relevant to biology they are easily rearranged by thermal motion. Proteins are consequently very dynamic molecules that are best understood as an ensemble of inter-converting conformers, rather than single static structures.
The majority of experimental techniques available for structural biology however, report on the ‘average’ positions of the constituent atoms. In order to fully understand how biological molecules function, it is necessary to move beyond static structures. NMR spectroscopy is unique in that it can simultaneously describe the motions that proteins undergo as well as their structure. It can therefore yield great insight into how proteins ‘work’.
There are two major types of experiments that we employ. The first, termed here as ‘fast’ measurements (above, green), are able to characterise motion on the pico/nano second timescale. In structured proteins, these relatively rapid motions include bond liberations, and side chain rotations. In unfolded proteins, these motions can be more extensive, reflecting the inter-conversion of many disordered conformations.
In addition to ‘fast’ measurements we probe slower motions (above, blue). Excitingly, it is becoming increasingly evident that even apparently immobile proteins often undergo infrequent, but substantial conformational rearrangements on the micro/mili second timescale, under biologically relevant solutions conditions. These higher energy conformations, termed ‘excited’ states, have been shown to play crucial functional roles in biochemical processes including molecular recognition, ligand binding, enzyme catalysis, cell signalling and protein folding, to name a few. We specialise in using ‘CPMG’ and ‘R1ρ’ NMR experiments to characterise these relatively slow dynamics. With these experiments, even if the population of an ‘excited’ state is on the order of 1% of the total (above, black), we can still characterise its structure and dynamics in almost the same detail as the majorly populated ‘ground’ state.
Solution NMR spectroscopy is often considered to have an inherent size limit of proteins only a few tens of kilo-Daltons in mass. However, exploiting the detailed spin physics and preparing carefully labelled samples, we are able to routinely perform both fast and slow dynamical measurements on proteins up to the mega-Dalton size range. To put this in context, without these improvements, approximately 10% of single chain proteins within the human genome are within the sights of solution NMR. With these improvements, 99% of the single chain proteins, and many of their complexes can potentially be analysed using solution NMR. When looking at complexes larger than 1MDa, we also employ solid state NMR techniques.
By putting together structural and dynamical information about the conformations that proteins populate using such ‘fast’ and ‘slow’ experiments, we can experimentally observe protein structure and dynamics in unprecedented detail.
Projects are available in both methods development, creating experiments that give us more information about ‘excited’ states, and in applications to a range of systems that are biochemically fascinating.