NMR was the first practical fragment-finding method, and
continues to be popular. Just over the past year we’ve discussed several new
techniques, (here, here, and here), and this post highlights three more.
In Angew. Chem. Int.
Ed., Jesus Angulo and colleagues at the University of East Anglia describe
differential epitope mapping by STD NMR (DEEP-STD NMR). STD NMR, the most
popular of ligand-detected methods according to our poll, can provide some information
as to which portions of a ligand are close to a protein, but doesn’t show where
on a protein the ligand binds. In DEEP-STD NMR, two separate NMR experiments
are conducted and the results compared to provide this information.
The researchers provide two implementation of the technique.
In the first, the protein is “irradiated” at two different frequencies; for
example, the aliphatic and aromatic regions. Protein residues that are directly
irradiated will show a stronger STD to ligand protons than those that are
indirectly irradiated, thus revealing whether one region of the ligand is
closer to an aromatic or an aliphatic amino acid side chain. If the structure
of the protein is known, this can then reveal the orientation of the ligand within
the binding site. A similar experiment can be done using H2O vs D2O
to determine whether a portion of a ligand is in close proximity to polar
residues in the protein.
Water is the subject of the second paper, in J. Med. Chem., by Robert Konrat and colleagues
at the University of Vienna and Boehringer Ingelheim. As we’ve previously
noted, water often plays a critical role in protein-ligand interactions. The
new method, called LOGSY titration, involves doing a series of WaterLOGSY
experiments at different protein concentrations and plotting the signals for
each proton in the ligand as a function of protein concentration; ligand
protons close to the protein show steeper slopes. The researchers examine pairs
of bromodomain ligands and demonstrate that LOGSY titration can confirm changes
in binding mode previously seen by crystallography. The technique could also
reveal what portions of the ligands make interactions with disordered water
molecules, which are more difficult to detect in crystal structures.
Both of these techniques provide useful but incomplete
information about ligand binding modes. A paper in J. Am. Chem. Soc. by Andreas Lingel and his Novartis colleagues
describes how to generate more detailed models. The researchers used a
deuterated protein in which all methyl groups (in methionine, isoleucine,
leucine, valine, alanine, and threonine) were 13C-labeled. Multiple intermolecular
NOEs between the protein and several previously characterized ligands were
collected and the resulting distances fed into modeling software to produce
good agreement with the known structures. More significantly, the researchers
were able to use the method prospectively with two weak (0.9 and 2.8 mM)
fragments. The binding models were sufficiently accurate to guide chemical
optimization, resulting in molecules with 30-50 µM affinities. Subsequent
crystal structures revealed that these bound as predicted. Impressively, this
was done on a protein that forms 115 kD hexamers – larger than those typically
tackled by NMR.
Teddy would normally close his NMR posts by stating –
usually quite forcefully – whether he felt the technique was practical or not. I’m
no NMR spectroscopist, so I’ll throw this question out to readers – do you plan
to try any of these approaches?
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