17 July 2017

Native mass spectrometry revisited

Native electrospray ionization mass spectrometry (ESI-MS) is one of the less-commonly used fragment finding methods. The technique relies on gently ionizing a protein-fragment complex without causing denaturation; bound fragments reveal themselves as shifts in mass. The technique is truly label-free, and can use very small amounts of protein and fragments. In practice the technique can work really well, reasonably well, or quite poorly. Two new papers shed light on factors that influence success.

The first paper, by Kevin Pagel (Freie Universität Berlin), Benno Kuropka (Bayer), and collaborators, examines four different cancer-related proteins. Let me say up-front that that the paper is remiss in not disclosing the chemical structures of any of the fragments, so in a very real sense this work is not reproducible. It is a shame the editors of ChemMedChem were not more demanding. That said, there is some useful information here.

Most of the focus is on the protein MTH1, screened at 10 µM concentration with 100 µM of each fragment. This was not a naïve screen; the fragments were previously identified from a thermal shift assay (TSA): 24 stabilized the protein, 4 destabilized it, and 5 had no effect. Remarkably, all of the fragments showed complexes in ESI-MS ranging between 6 – 66%, even those that had no effect in the TSA! Choosing an (admittedly arbitrary) 20% cutoff weeded out most of the false positives: 16 of the 24 stabilizers passed, while none of the destabilizers or neutral molecules did.

The best hit by ESI-MS also gave the strongest thermal shift, and a titration curve revealed an impressive dissociation constant of 1.7 µM. However, even at high concentrations of fragment the amount of bound complex did not exceed 70%, meaning that interpretation of single-dose experiments (for example, from a primary screen) could be problematic.

The results were similar for the protein KDM5B: 8 of 9 stabilizing fragments were hits by ESI-MS, as were two of 7 destabilizing fragments. Note that fragments that destabilize proteins can still be tight binders, as illustrated here.

For two additional proteins, however, ESI-MS was disappointing. For BRPF1, ESI-MS didn’t find any of the 11 hits from TSA, while for UHRF1 it found only a single hit – though this hit was not one of the 10 stabilizers identified by TSA. One could argue that the TSA hits were false positives were it not for the fact that, in the case of BRPF1, 6 of them were confirmed by crystallography.

The second paper, in Angew. Chem., comes from Chris Abell and coworkers at the University of Cambridge, and focuses on the protein EthR, a potential target for tuberculosis that we’ve previously discussed.

EthR binds to DNA, so rather than look for direct binding of fragments to EthR the researchers instead looked for fragments that could disrupt the EthR-DNA complex. A small library of 73 fragments was tested (at 0.5 mM each, in 2% DMSO), yielding 8 hits. The same library was screened under the same conditions using differential scanning fluorimetry (DSF), yielding 7 hits, 4 of which had also been identified using ESI-MS. All 11 of these molecules were then tested under the same conditions in an SPR assay to see if they could disrupt the interaction between EthR and chip-bound DNA. The 7 best SPR hits were all fragments that had been identified by ESI-MS. Moreover, two fragments – one identified solely by ESI-MS and one identified by both ESI-MS and DSF – were characterized bound to EthR crystallographically, and these represent new chemotypes for this target.

So what are we to make of all this? In common with other techniques, ESI-MS works well for some targets and less well for others. The problem is that it is not clear what distinguishes the two classes of targets. If you have access to the equipment and expertise you might consider adding ESI-MS to your screening cascade. But if you can only afford to buy one instrument for fragment screening, you’d probably be better off investing in NMR or SPR.

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