10 November 2019

A new tool for detecting aggregation

Historically the most popular method for finding fragments has been ligand-detected NMR. Preliminary results of our current poll (to the right) suggest crystallography has pulled ahead. (Please do vote if you haven’t already done so.) However, NMR has many uses beyond finding fragments, as illustrated in a recent J. Med. Chem. paper by Sacha Larda, Steven LaPlante, and colleagues at INRS-Centre Armand-Frappier Santé Biotechnologie, NMX, and Harvard.

Among the many artifacts that can occur in screening for small molecules, one of the most insidious is aggregation. A distrubing number of small molecules form aggregates in water, and these aggregates give false positives in multiple assays. Unfortunately, determining whether aggregation is occurring is not always straightforward. The new paper provides a simple NMR-based tool to do just that.

All molecules tumble in solution, but small fragment- or drug-sized molecules tumble more rapidly than large molecules such as proteins. The “relaxation” of proton resonances is faster in slower tumbling molecules, and in the NMR experiment called spin-spin relaxation Carr-Purcell-Meiboom-Gill (T2-CPMG) various delays are introduced and slower tumbling molecules show loss of resonances. Indeed, this technique has frequently been used in fragment screening: if a fragment binds to a protein, it will tumble more slowly, resulting in loss of signal.

The researchers recognized that an aggregate could behave like a large molecule, and they confirmed this to be the case for known aggregators, while non-aggregators did not. The experiment is relatively rapid (~30 seconds), and has been used to profile a 5000-compound library to remove aggregators.

One of the frustrations of aggregators is that it is currently impossible to predict whether a molecule will aggregate, and indeed, the researchers show several examples of closely related compounds in which one is an aggregator while the other is not. Even worse, the phenomenon can be buffer-dependent: the researchers show a fragment that aggregates in one buffer but not in another, even under the same pH.

Many fragment screens are done with pools of compounds, and the researchers find that molecules can show a “bad apple effect”, whereby previously well-behaved molecules appear to be recruited to aggregates.

The limit of detection for T2-CPMG is said to be single-digit micromolar concentration of small molecule, though the researchers note that double- or triple-digit micromolar concentrations are more practical, which is more typical of fragment screens anyway. And some compounds may show rapid relaxation due to non-pathological mechanisms, such as tautomerization or various conformational changes.

Still, this approach seems like a powerful means to rapidly assess hits, and pre-screening a library makes sense. Another NMR technique using interligand nuclear Overhauser effect (ILOE) has also been used to test for aggregation, though not to my knowledge so systematically. For the NMR folks out there, which methods do you think are best to weed out aggregators?

04 November 2019

Second harmonic generation (SHG) vs KRAS

Practical Fragments is currently running a poll on fragment-finding methods used by readers – please vote on the right-hand side. One biophysical method that perhaps we should have included is second harmonic generation (SHG). A recent paper in Proc. Nat. Acad. Sci. USA by Josh Salafsky, Frank McCormick, and collaborators at Biodesy, University of California San Francisco, and elsewhere describes the technique and its application to find fragments that bind to the oncogenic protein KRAS.

In SHG, two photons of the same energy are absorbed by a material which then emits a single photon with twice the energy. In the commercial instrument developed by Biodesy, a powerful 800 nm laser irradiates a dye, and the 400 nm photon it emits is detected. The intensity of the signal is exquisitely sensitive to the precise orientation of the dye. If a protein is labeled with an SHG-active dye and then immobilized on a glass surface, even subtle changes in conformation will be detected.

The researchers chose the G12D mutant form of KRAS, which is one of the most common variants and is associated with particularly aggressive tumors. They labeled the protein with a lysine-reactive SHG dye under conditions in which each protein would, on average, have one covalently-bound dye molecule (though some would have none and others would have more than one). Proteolysis and mass-spectrometry analysis revealed that the dye molecule labeled three different lysine residues, which the researchers viewed as a feature since a ligand causing a conformational change to any of the lysine residues would generate a signal. The researchers also demonstrated that the dye modification did not interfere with the ability of KRAS to bind to the RAS-binding domain of RAF.

Labeled KRAS was then immobilized and tested against several proteins known to bind it, including antibodies and the nucleotide exchange factor SOS. These produced SHG signals, presumably by causing conformational changes to KRAS, while non-binders such as tubulin did not.

Having established that the assay could detect binders, the researchers screened 2710 fragments at 250 and 500 ┬ÁM, and obtained a whopping 490 hits. These were then triaged by screening at lower concentrations and performing dose-titrations, and 60 were then characterized by SPR.

Fragment 18, 4-(cyclopent-2-en-1-yl)phenol, showed binding by both SHG and SPR, and was further studied by 2-dimensional NMR (1H-15N HSQC). This technique allowed measurement of the weak 3.3 mM dissociation constant. More importantly, it allowed the researchers to establish the binding location as being near the so-called “switch 2” region where SOS normally binds. This is the same region where a previous NMR screen had identified the slightly more potent fragment DCAI. The current paper confirmed that finding, though the researchers found evidence that DCAI may bind to other sites too. Docking studies using SILCS suggested that fragment 18 likely binds in a similar orientation as DCAI. Not surprising given the low affinity, the new fragment did not show functional activity in a biochemical screen.

SHG is an interesting approach, and the ability to rapidly assess protein conformational changes distinguishes it from other biophysical techniques. Site-specific labeling would produce more informative data on which regions of a protein move. However, I wonder if SHG is perhaps too sensitive, as evidenced by the large number of hits. Indeed, the researchers demonstrated that the promiscuous lipophilic amine mepazine also generated a strong SHG signal with KRAS. It would be interesting to do a head-to-head comparison with other similarly rapid techniques such as DSF or MST. Have you tried using SHG, and if so, how did it perform for you?