Labeled ligand displacement: Extending NMR-based screening of protein targets

Article abstract of DOI:10.1021/ml1000849

NMR spectroscopy has enjoyed widespread success as a method for screening protein targets, especially in the area of fragment-based drug discovery. However, current methods for NMR-based screening all suffer certain limitations. Two-dimensional methods like SAR by NMR require isotopically labeled protein and are limited to proteins less than about 50 kDa. For one-dimensional, ligand-based methods, results can be confounded by nonspecific compound binding, resonance overlap, or the need for a special NMR probe. We present here a ligand-based method that relies on the exchange broadening observed for a ¹³C-labeled molecule upon binding to a protein target (labeled ligand displacement). This method can be used to screen both individual compounds and mixtures and is free of the artifacts inherent in other ligand-based methods.

Full text of DOI:10.1021/ml1000849

TECHNOLOGY NOTE
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Labeled Ligand Displacement: Extending NMR-Based
Screening of Protein Targets
Steven L. Swann, Danying Song, Chaohong Sun, Philip J. Hajduk, and Andrew M. Petros*
Global Pharmaceutical Research and Development, Abbott Laboratories, 100 Abbott Park Road, Abbott Park, Illinois 60064
ABSTRACT NMR spectroscopy has enjoyed widespread success as a method for
screening protein targets, especially in the area of fragment-based drug discovery.
However, current methods for NMR-based screening all suffer certain limitations.
Two-dimensional methods like “SAR by NMR” require isotopically labeled protein
and are limited to proteins less than about 50 kDa. For one-dimensional, ligand-
based methods, results can be confounded by nonspecific compound binding,
resonance overlap, or the need for a special NMR probe. We present here a ligand-
based method that relies on the exchange broadening observed for a 13C-labeled
molecule upon binding to a protein target (labeled ligand displacement). This
method can be used to screen both individual compounds and mixtures and is free
of the artifacts inherent in other ligand-based methods.
KEYWORDS NMR, screening, fragment, labeling, ligand
ver the last few decades, nuclear magnetic reso-
nance (NMR) spectroscopy has developed into a
powerful tool for the study of biological molecules
molecules. While the approach continues to be the most robust
and reliable method for NMR-based screening of protein targets,
it suffers from several disadvantages. One major limitation of
traditional SAR by NMR is protein size. As the size of a protein
increases, its rotational correlation time decreases, and subse-
quently, the resonance line width increases. The heteronuclear
single quantum coherence (HSQC) spectrum upon which this
method is based not only becomes more crowded due to a
greater number of resonances, but the individual resonances
become broader until the spectrum is no longer interpretable. In
using this method, the realistic upper limit for the protein
molecular mass is roughly 50 kDa. In addition, protein aggrega-
tion can rapidly degrade the quality of the HSQC spectrum
regardless of protein size and can be a major hindrance to the
SAR by NMR method. Finally, SAR by NMR requires isotopically
labeled (either 15N or 13C) protein, which, in many cases, can be
difficult to obtain.
In light of these limitations, other NMR-based methods have
been developed, which, by contrast, monitor the change of a
specific ligand property upon binding to a target protein. These
ligand-based methods include saturation transfer difference
(STD) spectroscopy,6 waterLOGSY,7 SLAPSTIC,8 TINS,9 FAXS,10
one-dimensional direct competition methods,11 and others.3
Unfortunately, ligand-based methods also suffer from various
limitations, the most common of which are interference from
nonspecific binding of a small molecule to a target protein and
resonance overlap in the one-dimensional spectra that are
typically recorded for these ligand-based methods. The excep-
tion to this is the fluorine-based FAXS method. Therefore, there
is a need for an improved and more reliable ligand-based
O
and systems.1 The structure determination of small, well-
behaved proteins and polynucleotides in solution using NMR
has become routine. To complement this structural informa-
tion, NMR spectroscopy can also provide information on the
molecular dynamics of these macromolecules.2 However, its
most important contribution, especially in the area of phar-
maceutical research, may be the ability of NMR spectroscopy
to provide information about molecular interactions at the
atomic level. This information includes, first and foremost,
determining whether or not a particular small molecule
binds to a protein of interest. Second, if the molecule does
bind, which atoms of both the small molecule and the
protein are involved in this interaction? Various NMR-based
techniques have been developed to address these two
points. These methods provide varying levels of detail and
have been met with varying degrees of success.3
One of the earliest of these techniques to be developed
was “structure-activity relationships (SAR) by NMR”, which
was the first method to be truly applied in a screening mode,
that is, testing thousands of small molecules for binding to a
protein target.4 Since the initial publication describing SAR
by NMR, the use of NMR in lead discovery and optimization
has become widespread in the pharmaceutical industry.
Most notably, SAR by NMR played an integral role in the
discovery of the pro-apoptotic molecules ABT-737 and ABT-
263, the latter of which is currently being tested clinically as
an anticancer agent.5
SAR by NMR is a target-based method where the chemical
shift of protein resonances, from either backbone amides or
side chain methyl groups, is monitored as the protein is
interrogated with a single small molecule or a mixture of small
Received Date: April 22, 2010
Accepted Date: June 15, 2010
Published on Web Date: June 24, 2010
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Figure 2. Effect of increasing protein (Hsp90, ATPase domain)
concentration on the aromatic resonances of compound 1 (20 uM)
in a proton NMR spectrum. (A) No protein, (B) 5 μM protein, (C)
10 μM protein, and (D) 30 μM protein.
between free and bound species. For a ligand that is in the
fast to intermediate exchange regime with a protein target,
the exchange-broadening contribution to the resonance line
width at half height is given by:
- 1
Δv1=2 ¼ ðτAπÞ
where τA is the protein-bound lifetime of the ligand. Two
factors affecting this bound lifetime are the inherent affinity
of the ligand for the protein, which can be expressed as a
dissociation constant, KD (equal to the ratio of kon to koff), and
the ratio of the protein to the ligand in solution. Thus, for a
ligand with a given affinity for its protein target, there is a
ratio of protein to ligand, which can be determined empiri-
cally, wherethis exchange-broadening contribution will be at
its maximum.
This exchange-broadening effect could, in principle, be em-
ployed either directly or in competition mode, much like STD, to
monitor ligand binding to a protein. To directly observe binding
of a ligand to a protein, the ligand must have at least one
resonance that does not overlap with signals from the added
protein. However, for most ligands, this is most likely the
exception rather than the rule. Similarly, to be used in a
competition mode, the “probe” molecule must have at least
one resonance that does not overlap with signals from the
protein or with signals from potential binders. This overlap
would likely preclude the screening of mixtures.
This dilemma is illustrated in Figure 2 for the binding of
compound 1 (Figure 1) to the ATPase domain of heat-shock
protein 90 (Hsp90).12 Hsp90 is an intracellular molecular
chaperone protein that consists of three domains and is widely
expressed in various cells.13 Targeting of the amino-terminal
ATPase domain (MW ∼ 28 kDa) in cancer cells by various
molecules has been shown to have an antiproliferative effect.14
Figure 2A shows the aromatic region of a one-dimensional pro-
ton spectrum of compound 1 in solution. This compound binds
to the ATPase domain of Hsp90 with an inhibition constant (Ki)
of 14 μM as determined in a fluorescence resonance energy
transfer assay (TR-FRET).12 As shown in Figure 2B-D, the
addition of increasing amounts of protein causes broadening of
the ligand signals. However, it is virtually impossible to monitor
Figure 1. Various compounds employed in this study.
method that overcomes these limitations and can be executed
using standard triple-resonance technology.
Herein, we describe a new ligand-based method that we
have termed “labeled ligand displacement” (LLD), which is
based on the displacement of an isotopically (13C) labeled
ligand from a protein target of interest upon addition of an
individual compound or a mixture of compounds. The
method is not limited by the size of the protein, is less
sensitive to nonspecific compound binding than other li-
gand-based methods, and does not require preparation of
isotopically labeled protein. It compares favorably to STD,
the most widely employed ligand-based method, and offers
some distinct advantages. Finally, LLD can be implemented
with standard NMR technologies, including standard triple-
resonance probes, and will greatly increase the number of
protein targets for which NMR-based screening, especially
fragment screening, can be applied.
The interaction of most small ligands with a protein target
can be treated in terms of a simple two-site exchange
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Figure 4. (A-D) Effect of increasing protein (carbonic anhydrase)
concentration on the 13C-selected, proton NMR spectrum of
compound 4 (20 μM). (A) No protein, (B) 10 μM carbonic anhy-
drase, (C) 20 μM carbonic anhydrase, and (D) 40 μM carbonic
anhydrase. (E-G) LLD spectra for compound 4 (20 μM) in the
presence of 40 uM protein plus increasing concentrations of
compound 5: (E) 100 μM, (F) 200 μM, and (G) 400 μM.
Figure 3. (A-D) Effect of increasing protein (Hsp90, ATPase
domain) concentration on the aromatic resonances of compound
2 (20 μM) in a 13C-selected, proton NMR spectrum. (A) No protein,
(B) 10 μM protein, (C) 20 μM protein, and (D) 30 μM protein. (E-G)
LLD spectra for compound 2 (20 μM) in the presence of 30 μM
protein plus increasing concentrations of compound 3: (E) 100 μM,
(F) 200 μM, and (G) 400 μM.
this broadening at higher protein concentrations due to reso-
nance overlap with signals from the protein itself.
To overcome this limitation, a version of compound 1,
which contains a uniformly 13C-labeled phenyl ring, was
prepared by coupling 4-chloro-6-methylpyrimidin-2-amine
with 13C6-labeled phenyl boronic acid (Supporting In-
formation). Figure 3A shows a one-dimensional 13C-se-
lected proton spectrum of the 13C-labeled ligand in buffer
alone. This spectrum is essentially the first slice of a 13C-
HSQC spectrum. In panels B-D is shown the effect of
increasing concentrations of the unlabeled ATPase domain
of Hsp90 on the ligand resonances. In contrast to the case
with unlabeled ligand, it can clearly be seen that at a 1:1
molar ratio of protein to ligand, the ligand resonances are
completely broadened out (Figure 3C). Upon addition of a
competing ligand (compound 3, Ki ∼ 11 μM), signals from
the labeled probe are recovered in a dose-dependent
manner (Figure 3E-G). In spectra F and G, where com-
pound 3 is at 200 and 400 μM, respectively, additional
small signals appear at about 7.5 and 8.0 ppm from the
1.1% natural abundance of 13C in compound 3.
A further illustration of the utility of isotopic labeling to
cleanly monitor exchange broadening is shown for the bind-
ing of 13C-labeled 4-fluorobenzenesulonamide (compound 4,
Figure 1) to bovine carbonic anhydrase. Carbonic anhydrases
are ubiquitous enzymes that catalyze the conversion of carbon
dioxide to bicarbonate ion and play an important role in various
physiological processes including respiration and certain biosyn-
thetic reactions.15 Bovine carbonic anhydrase is a ∼30 kDa
protein that is inhibited by various sulfonamide-based ligands.
Using surface plasmon resonance, the affinity of an unlabeled
version of compound 4 was found to be about 1 μM for bovine
carbonic anhydrase.16 Compound 4 was made by treating
uniformly 13C-labeled 4-fluorosulfonylchloride with ammonia
in methanol (Supporting Information). Figure 4 illustrates
the reduction in intensity of the aromatic resonances from the
ligand in response to increasing protein concentration. At a 2:1
molar ratio of protein to ligand, the resonances from the free
Figure 5. LLD method applied to screening mixtures. (A) Reference
spectrum of Hsp90, ATPase domain plus probe (30 μM protein and
20 μM compound 2). (B) Plus mixture of nine compounds (10-18),
at 200 μM each, which do not bind to the protein. (C) Addition of
200 μM compound 3 (Ki ∼ 6 uM). (D) Probe alone (20 μM).
ligand have completely broadened out. The two broad signals in
spectrum D presumably arise from bound ligand. As shown for
the Hsp90, ATPase domain, the addition of a competing ligand
(compound 5, Ki ∼ 48 uM) results in recovery of the probe
signals. In addition to utilizing specifically designed and labeled
small molecules as probes, the broadening effect can also be
observed for the binding of natural products, such as nucleotides
and peptides, to proteins (Supporting Information, Figures 1S
and 2S).17,18
Besides the screening of individual compounds, LLD can
be used to screen mixtures of compounds against a protein
target. Figure 5A shows a spectrum of the Hsp90, ATPase
domain plus probe (compound 2). In spectrum B, a mixture
of nine compounds (10-18 in Figure 1), which do not bind to
the protein, has been added. These ligands show no pertur-
bation of the protein 13C-methyl resonances in a 13C-HSQC
spectrum when added at an approximately 10-fold molar
excess. As can be seen, no recovery of the probe signals
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Resonance overlap is also a potential drawback of the one-
dimensional competition method described by Siriwardena
et al.11 In this method, the intensity of an unlabeled probe
molecule in the presence of its protein target is monitored as
potential binders are added. As with STD, a probe molecule
must be found whose NMR signals do not overlap with those
from the probe, a limitation that is overcome with the LLD
method.
In choosing the 13C-labeled ligand (probe) for an LLD experi-
ment, two factors need to be considered. The first is the affinity
of the probe for the protein. In our experience, probe molecules
should ideally have an affinity of between 1 and 100 μM. For
probes that bind more weakly, a large excess of protein may
need to be added to observe sufficient exchange broadening. In
contrast, it may be difficult to compete away a probe molecule
that binds too tightly to its protein target, especially with weakly
binding fragments. The second factor to consider is the ease
with which an isotopically labeled probe molecule can be
obtained. Isotopically labeled amino acids, peptides, and nu-
cleotides are, in general, easily obtained from commercial
sources. Synthetic ligands present more of a challenge, as the
appropriate isotopically labeled starting materials must be
found. However, because only part of a synthetic ligand needs
to be labeled, that is, a single phenyl moiety or methyl group, in
order for the molecule to work as a probe, this aspect of the
method is not as limiting as it may appear.
Screening a protein target for potential binders is carried out
by first preparing a stock solution of the isotopically labeled
probe and unlabeled protein at the appropriate, experimen-
tally determined, protein to ligand ratio. Compounds can then
be added to this stock solution individually or as mixtures.
Compound binding will result in displacement of the probe
and subsequent recovery of signal. For a series of ligands
added at the same concentration, the amount of signal reco-
vered will be proportional to the affinity of each ligand for the
protein (Supporting Information, Tables 1 and 2). The sensi-
tivity of this method as a screening tool can be adjusted in two
ways, either by varying the concentration of potentially com-
peting ligands or by varying the concentration of the target
protein. Higher ligand concentrations can be used to increase
the sensitivity of the method for screening weakly binding
fragments, while lower concentrations can be used to increase
the selectivity of the method. Analogously, the sensitivity/
selectivity of the method can be adjusted by varying the
amount of target protein. The addition of protein above the
amount needed to observe maximal broadening will increase
the selectivity of the method as the additional protein will act
as a “sink” for competing ligands.
Figure 6. STD-competition spectra for the Hsp90, ATPase domain.
(A) STD spectrum of probe molecule (compound 1) at 100 μM in the
presence of 10 μM protein, (B) plus 100 μM compound 3, (C) plus
200 μM compound 3, and (D) plus 400 μM compound 3.
is observed. The broad background signals that appear
between 7.2 and 8.0 ppm are from the 1.1% natural
abundance of 13C in these ligands, which are each at 200
μM. Upon addition of 200 μM compound 6 (Ki ∼ 5 μM),
enough probe signal is recovered above the background that
this compound could clearly be flagged as a binder.
Of all of the ligand-based screening methods using NMR,
STD spectroscopy continues to be the most broadly applied
ligand-based method. It has been applied in both direct and
competitive modes and has been used not only to identify
protein ligands but also in mapping the binding epitopes of
protein-bound ligands.6,19 The basis for the STD-NMR ex-
periment is the transfer of magnetization from a protein to a
bound ligand. While it is a very simple and versatile tool for
analyzing protein-ligand interactions, it does suffer from a
few significant drawbacks, among which are interference
from nonspecific binding and signal overlap.
Figure 6 illustrates the LLD competition experiment of
Figure 3E-G but carried out in STD mode. In Figure 6A is the
aromatic region of the STD spectrum for compound 1 bound
to the Hsp90, ATPase domain. The ligand to protein ratio in this
case is 10:1, which is typical for an STD competition experi-
ment. Spectra B-D show the effect of adding increasing
amounts of a competing ligand, compound 3. With compound
3 at 100 μM (spectrum B), there is a clear decrease in the STD
signal. However, as additional compound is added, there is no
further decrease in the STD signal (spectra C and D). We have
observed this with several other systems in our lab and have
attributed it to nonspecific interaction of the STD probe mole-
cule with the protein. This nonspecific binding gives rise to a
certain “background” STD signal, which remains even upon
further addition of competing ligand. Figure 6 also illustrates the
second drawback of the STD-competition method, which is
the potential overlap of signals from a competing ligand with
the signals from the STD probe molecule. In this particular case,
there is only a slight overlap of a resonance from added
compound 3 (∼7.59 ppm) with a resonance from the probe.
However, it is clear from this figure how resonance overlap
could be a serious drawback, especially for screening mixtures
of compounds. Ideally, one can choose a probe with a reso-
nance well-removed from signals of potential binders, but this
may not always be possible.
In addition to simple binding conformation, this method
could be used to quickly obtain a rough estimate of the KD
value fora given ligand, provided that a few ligands of known
and varying affinity are available. These known ligands could
be used to derive a calibration curve by plotting the percent
recovered intensity for each ligand against its affinity. Re-
covered intensity values for ligands of unknown affinity
could then be estimated from this curve.
We have developed a new ligand-based method for NMR
screening that we have named LLD as it is based on the
displacement of a carbon-13-labeled probe molecule from a
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protein target. It is a very simple method to apply and has
advantages over both the traditional protein-based screen-
ing methods such as “SAR by NMR” as well as other ligand-
based methods such as STD. As compared to protein-based
methods, LLD can be applied to any size protein, and in
fact, the broadening effect on which the method is based
should be enhanced for larger proteins. While the amounts
of protein needed are about the same as for protein-
based methods, isotopically labeled protein is not needed,
and thus, protein from sources other than recombinant
Escherichia coli grown in minimal media can be used. As
compared to STD methods, LLD allows one to filter out
signals from competing ligands, which is especially advanta-
geous for the screening of mixtures. In addition to the utility
of LLD for traditional small molecule or fragment screening,
it could also, in principle, be used as an orthogonal assay for
essentially any soluble protein target.
affinity to proteins via magnetization transfer from bulk
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SUPPORTING INFORMATION AVAILABLE Experimental
procedures along with two figures showing the exchange-broad-
ening effect for a 13C-labeled peptide bound to the protein Bcl-xL
and for 13C-labeled ATP bound to Hsp90 and two tables that show
the amount of signal recovered upon addition of competing ligands
for both Hsp90 and carbonic anhydrase. This material is available
free of charge via the Internet at http://pubs.acs.org.
AUTHOR INFORMATION
Corresponding Author: *To whom correspondence should be
addressed. E-mail: andrew.petros@abbott.com.
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cations for inhibitors and activators. Nat. Rev. Drug Discovery
2008, 7, 168–181.
ACKNOWLEDGMENT Thanks to Sally Dorwin for preparation
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inhibitor interaction. J. Biomol. Tech. 2003, 14, 247–269.
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J. E.; Eberstadt, M.; Yoon, H. S.; Shuker, S. B.; Chang, B. S.;
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