ACS Medicinal Chemistry Letters
Letter
absence of binding (true negative) or limitations of the screening
setup at detecting weak binding (false negative).
To extend the binding detection range, we set to modify the
NMR experimental conditions by first increasing the protein
concentration to 40 μM while maintaining ligand concentrations
fixed at 1 mM (setup 2); and second by increasing concentrations
of protein and ligand to 30 μM and 3 mM (set-up 3), res-
pectively, thereby maintaining a protein/ligand excess of 100-
fold as in set-up 1.
We first applied the revised set-ups to the capped hydroxypro-
line (Hyp) core fragment 6, that had successfully yielded an
X-ray bound structure17 but had otherwise proven elusive to
biophysical detection. Each NMR experiment distinctly detected
binding of 6 under both set-ups 2 and 3 but not set-up 1, placing
this compound as a true positive hit (Figures 3 and S6,
Supporting Information).
Figure 1. Structural representation of the VCB multiprotein complex
and the pVHL:HIF-1α interface.
Set-up 2 provided the most reliable detection profile for 6
in CPMG and WaterLOGSY, a direct result of increasing the
fractional bound ligand, while maintaining total ligand
concentration constant at 1 mM. In contrast, set-up 3 gave the
best result in STD, as this technique is unaffected by increasing
free ligand concentration. Similarly, we were able to un-
ambiguously detect binding of compounds 8−11 to VCB,
whereas binding of compounds 7 and 12 remained undetected
under each revised setup (Table 1 and Figures S7 and S12,
Supporting Information). This highlighted that the newly
adopted set-ups enable robust discrimination between true
binders and nonbinders, which is a critical requisite in biophysical
fragment screening.
Aiming further characterization of the rescued binders, we
asked if they targeted specifically the pVHL-HIF-1α interface
and whether they would recapitulate the binding mode shown
as part of the intact parent compounds 1 and 2. To address this,
we first attempted to compete binding of 6 and 8−11 using a
high-affinity 19-mer HIF-1α peptide. Compounds 6, 8, and 11
were displaced by the peptide, placing them at this PPI (Table 1
and Figures S6, S8, and S11, panels a, b and c, Supporting
Information). To assess binding affinity for the displaced
compounds, competitive ITC experiments were carried out
using inhibitor 2 as the titrant in the presence of fragments 8
and 11, yielding apparent Kd of 2.7 and 4.3 mM for 8 and 11,
respectively (Table 1 and Figure S8 and S11, panel g, Supporting
Information). A matching Kd of ∼5 mM was obtained for 6 under
both direct and competitive conditions, thereby validating the
approach (Figure S6, panels g and h, Supporting Information).
These fragments maintained similar LE values (Table 1) of
the parent inhibitors, which notably fell around the value of
0.24 kcal mol−1 NHA−1 obeying LE’s generally observed for
PPI-targeting small molecules.1
should be <3).17 Moreover, a screen of ∼1300 Ro3 obeying
fragments library proved unsuccessful for targeting the
pVHL:HIF-1α interface.17 We were intrigued by these observa-
tions, in part because there is a growing belief that target
druggability correlates with hit rates from fragment screening.11
To resolve this conundrum, we decided to investigate the ability
to triage bona fide weak binding fragments using ligand-based
NMR spectroscopy, arguably one of the most sensitive
biophysical techniques that is widely applied to hit generation
in drug discovery.21
A library of 12 compounds was designed by defragment-
ing known inhibitors 1 and 2 (see Figures S1 and S3,
Supporting Information),18,19 and screened against the target
1
protein using three distinct ligand-based 1D H NMR experi-
ments (Table 1): first, saturation transfer difference (STD)22
experiments apply a selective pulse to saturate protein
resonances. Only ligands that bind to the protein will receive
saturation transfer, resulting in their signals to appear as positive
in a difference spectrum between the unsaturated and saturated
spectra; second, Carr−Purcell−Meiboom−Gill (CPMG)23
experiments exploit the faster T2 relaxation times of macro-
molecules relative to small molecules. Upon binding to the
protein, ligands relaxation time will decrease, causing a line
broadening and a consequent decrease in intensity on their
signals; finally, the water-ligand observed via gradient spectros-
copy (WaterLOGSY)24 experiments apply a selective pulse to
saturate resonances of water molecules. In the absence of protein,
cross-relaxation from water will yield control ligand signals that
are phased downward; in the presence of protein, small molecule
binders will receive an NOE contribution from water that is of
opposite sign relative to control, resulting in their signals either
showing a reduction in intensity, or even pointing upward.
Experimental conditions of 1 mM ligand and 10 μM protein,
which are typically used in NMR fragment screening, enabled
unambiguous detection in each of the three NMR experiments of
the larger, Ro3-breaking compounds 3−5 (set-up 1, Table 1).
We were also able to characterize binding thermodynamics by
isothermal titration calorimetry (ITC) and determine bound
X-ray structures by crystallographic soaks, confirming the
expected binding modes at the HIF-1α site (Table 1, Figures 2
and S4−S5, Supporting Information). In contrast, binding
detection was unfruitful for Ro3-compliant fragments 6−12
under set-up 1 (Table 1 and Figures S6−12a−c, Supporting
Information). This observation is consistent with our pre-
vious results,17 raising the question whether this is due to the
To gain information about the fragments binding mode, we
first turned to group epitope mapping (GEM) characterization
of their STD-NMR spectra.25 Relative degrees of saturation of
the individual protons were normalized to the highest saturated
proton of the compounds, yielding some information of the
proximity of each proton and the interacting protein surface.
The NMR GEM data for compound 6 (Table S2, Supporting
Information, and Figure 4) suggests the protons adjacent to the
hydroxyl group to be in closer contact with the protein, in perfect
agreement with the binding mode observed in the X-ray structure
(Figure S6, panel i, Supporting Information). In contrast, the
GEM data for 8 and 11 (Tables S3−4, Supporting Information)
could not conclusively inform about their binding modes as
24
dx.doi.org/10.1021/ml400296c | ACS Med. Chem. Lett. 2014, 5, 23−28