DOTA-amide lanthanide tag for reliable generation of pseudocontact shifts in protein NMR spectra

Article abstract of DOI:10.1021/bc200353c

Structural studies of proteins and protein-ligand complexes by nuclear magnetic resonance (NMR) spectroscopy can be greatly enhanced by site-specific attachment of lanthanide ions to create paramagnetic centers. In particular, pseudocontact shifts (PCS) g

Full text of DOI:10.1021/bc200353c

ARTICLE
pubs.acs.org/bc
DOTA-Amide Lanthanide Tag for Reliable Generation of
Pseudocontact Shifts in Protein NMR Spectra
†
§
†
†
†
§
Bim Graham, Choy Theng Loh, James David Swarbrick, Phuc Ung, James Shin, Hiromasa Yagi,
§
†
†
‡
§
Xinying Jia, Sandeep Chhabra, Nicholas Barlow, Guido Pintacuda, Thomas Huber, and
,§
Gottfried Otting*
§
Research School of Chemistry, Australian National University, Canberra ACT 0200, Australia
†
Medicinal Chemistry and Drug Action, Monash Institute of Pharmaceutical Sciences, Parkville VIC 3052, Australia
‡
Centre de RMN ꢀa Tr ꢀe s Hauts Champs, ENS-Lyon, 69100 Villeurbanne, France
S Supporting Information
b
ABSTRACT: Structural studies of proteins and proteinꢀligand
complexes by nuclear magnetic resonance (NMR) spectroscopy
can be greatly enhanced by site-specific attachment of lanthanide
ions to create paramagnetic centers. In particular, pseudocontact
shifts (PCS) generated by paramagnetic lanthanides contain
important and unique long-range structure information. Here,
we present a high-affinity lanthanide binding tag that can be
attached to single cysteine residues of proteins. The new tag has
many advantageous features that are not available in this
combination from previously published tags: (i) it binds lantha-
nide ions very tightly, minimizing the generation of nonspecific
effects, (ii) it produces PCSs with high reliability as its bulkiness
prevents complete motional averaging of PCSs, (iii) it can be
attached to single cysteine residues, alleviating the need of detailed prior knowledge of the 3D structure of the target protein, and (iv)
it does not display conformational exchange phenomena that would increase the number of signals in the NMR spectrum. The
performance of the tag is demonstrated with the N-terminal domain of the E. coli arginine repressor and the A28C mutant of human
ubiquitin.
’
INTRODUCTION
nature of lanthanide-induced PCSs makes them powerful tools
4,5
for structure determinations of proteinꢀprotein and proteinꢀ
ligand complexes (for recent reviews, see refs 6 and 7). In
practice, eight parameters suffice to fit Δχ tensors to the 3D
structure of a protein (x, y, z coordinates of the metal ion, three
Euler angles to relate the orientation of the Δχ tensor to the
frame of the protein atom coordinates, and the Δχ and Δχ
Site-specific attachment of paramagnetic lanthanide ions to
proteins produces large effects in NMR spectra that contain
1,2
valuable long-range structural information. Among the para-
magnetic effects, pseudocontact shifts (PCS) stand out for their
ease of measurement (as the difference in chemical shifts
observed for samples labeled with, respectively, a paramagnetic
or a diamagnetic lanthanide ion) and high information content
ax
rh
parameters). Lanthanides are uniquely suited for PCS measure-
ments, as different lanthanides produce different magnitudes of
Δχ tensors while their chemical properties are very similar. This
avoids chemical shift changes due to chemical differences among
(
as PCSs can be observed for nuclear spins beyond 40 Å from the
metal ion).
3The PCS, Δδ
PCS
(in ppm), of a nuclear spin can be described
3+
3+
3+
different paramagnetic and diamagnetic (La , Y , Lu ) ions.
A lanthanide ion can generate large PCSs only if it is site-
specifically attached to the target protein in a rigid manner. If the
tether between the lanthanide and the protein is flexible, the Δχ
tensor associated with the lanthanide changes its orientation
relative to the protein and the resulting averaging over positive
and negative PCS values greatly reduces the magnitude of
by
ꢀ
ꢁ
ð1Þ
1
2πr3
3
PCS
2
2
Δδ
¼ 1
Δχ ð3 cos θ ꢀ 1Þ þ Δχ sin θ cos 2j
ax
rh
2
where Δχ and Δχ denote, respectively, the axial and rhombic
ax
rh
components of the magnetic susceptibility anisotropy (Δχ)
tensor and r, and θ and j are the polar coordinates of the
nuclear spin with respect to the principal axes of the Δχ tensor.
The limited number of parameters in eq 1 and the long-range
Received: July 5, 2011
Revised:
August 9, 2011
Published: August 31, 2011
r 2011 American Chemical Society
2118
dx.doi.org/10.1021/bc200353c
|Bioconjugate Chem. 2011, 22, 2118–2125

Bioconjugate Chemistry
ARTICLE
observable PCSs. Therefore, no PCS may be observed if the
tether allows substantial reorientation of the lanthanide ion with
respect to the protein.
via syringe to a solution of (S)-1-phenylethanamine (15.3 g,
126 mmol) inDCM (200 mL) at0 °C under nitrogen. Stirringwas
continued for a further 2 h at room temperature. The solution was
washed with 2 N HCl (100 mL) followed by saturated brine
The problem of rigid lanthanide attachment can be addressed
in different ways. Lanthanide chelates with two arms for attach-
ment to two cysteine residues have been shown to minimize
(100 mL), then dried (MgSO ), and solvent removed under
4
1
reduced pressure to afford 1 (12.7 g, 83%) as a white solid. H
8
ꢀ10
residual motion of the lanthanide relative to the protein.
A
NMR (400 MHz, CDCl ) δ 7.40ꢀ7.25 (m, 5H), 6.79 (br s, 1H),
3
related strategy combines an N-terminal fusion with a lanthanide
binding peptide (LBP) with a disulfide bond between the LBP
5.12 (apparent p, J = 7.0 Hz, 1H), 3.95ꢀ3.82 (m, 2H), 1.55 (d,
13
J = 6.9 Hz, 3H). C NMR (101 MHz, CDCl ) δ 164.7, 142.5,
3
5
,11
and the target protein.
between two strands of a β-sheet similarly immobilizes
Engineering an LBP into the turn
128.9, 127.7, 126.2, 49.7, 29.4, 21.7. LRMS (ESI) m/z 244
8
1
+
79
+
(7%, [M[ Br]H] ), 242 (7, [M[ Br]H] ), 138 (10), 136 (9),
105 (100).
1
2
lanthanides. Another strategy coordinates the lanthanide by
groups from two different sites of the protein. One group may be
a small lanthanide binding tag attached to a cysteine residue,
while the other group can be a carboxyl group of the protein.
Small tag molecules can also be attached to two cysteines with the
0
00
2,2 ,2 -(1,4,7,10-Tetraazacyclododecane-1,4,7-triyl)tris-
(N-((S)-1-phenylethyl)acetamide) (2). A solution of 1 (18.4 g,
76 mmol) in chloroform (500 mL) was added dropwise to a
stirred mixture of cyclen (4.37 g, 25 mmol) and DIPEA (35 mL,
200 mmol) in chloroform (500 mL) under a nitrogen atmo-
sphere at room temperature. The mixture was stirred for a further
24 h, then washed with water, and the organic phase dried over
13,14
15,16
aim of coordinating a single lanthanide between the two tags.
Finally, a lanthanide binding site can be engineered into the
target protein by introducing amino acids with negatively
1
7
charged side chains. All these strategies rely on prior knowledge
of the 3D structure of the target protein, at least at the site of
modification.
MgSO . After removal of the solvent, the residue was purified by
4
flash chromatography (gradient from 90:10:1 to 80:20:2 EtOAc/
MeOH/NH ) to afford the product (10.2 g, 61%) as a thick, light
3
1
To harness the power of PCSs for 3D structure determina-
tions, however, it is necessary to use lanthanide tags that can be
site-specifically attached without detailed knowledge of the 3D
structure of the target protein. In this situation, tags with single
attachment points have an advantage even if they may only
incompletely immobilize the lanthanide ion. For example, LBPs
with single cysteine residues have been shown to deliver large
PCSs following attachment to the target protein via a disulfide
yellow oil. H NMR (400 MHz, DMSO) δ 8.36 (d, J = 8.0 Hz,
2H), 8.26 (d, J = 7.8 Hz, 1H), 7.33ꢀ7.26 (m, 12H), 7.25ꢀ7.16
(m, 3H), 4.98ꢀ4.85 (m, 3H), 3.15ꢀ3.00 (m, 6H), 2.73ꢀ2.43 (m,
13
16H), 1.41ꢀ1.30 (m, 9H). C NMR (101 MHz, DMSO) δ
169.7(2), 169.6(8), 144.7, 144.4, 128.2, 128.2, 126.6, 126.5, 126.0,
126.0, 57.7, 55.9, 52.7, 52.3, 50.9, 47.7, 47.6, 46.4, 22.5, 22.4. LRMS
+
2+
(ESI) m/z 656 (85%, [M+H] ), 329 (100, [M+2H] ).
2-Chloro-N-(2-(pyridin-2-yldisulfanyl)ethyl)acetamide (3).
To a stirred solution of 2-(pyridin-2-yldisulfanyl)ethanamine
hydrochloride (3.00 g, 13.5 mmol) and chloroacetic acid (1.28 g,
14 mmol) in 1:10 dichloromethane/acetonitrile (50 mL) was
added DIPEA (7.0 mL, 400 mmol) and BOP (6.63 g, 15 mmol).
The reaction mixture was stirred overnight at room temperature
under nitrogen. Upon completion of the reaction, the solvent was
removed under reduced pressure and the resulting crude product
1
8,19
bond.
The bulkiness of the tag restricts the amplitude of
motion relative to the protein by steric constraints, preventing
excessive averaging of the PCSs. Similarly, tags based on lanthanide
complexes derived from cyclen (1,4,7,10-tetraazacyclododecane)
can deliver PCSs of useful magnitudes, while providing excep-
tionally small dissociation constants of the lanthanide from the
tag and robustness against extreme conditions of pH, tempera-
8,20
ture, and denaturants. As a drawback, synthesis of cyclen tags
can be expensive and arduous to prepare, and conformational
was dissolved in saturated NaHCO solution (150 mL) and
3
washed with ether (3 ꢁ 100 mL). The combined organic layers
20
equilibria can result in multiple sets of NMR signals.
were washed with saturated brine, dried (MgSO ), and the
4
Lanthanide complexes of DOTA derivatives display two
solvent removed under reduced pressure. Flash chromatography
21
fundamental conformational equilibria. The first affects the
chirality of the cyclen ring. The second equilibrium affects the
chirality of metal coordination by the acetate groups. Both
equilibria are coupled. It has been shown that the equilibria
can be pushed toward a single conformation of the lanthanide
complex by incorporating chiral centers in the cyclene ring and
(30:70 ethyl acetate/petroleum spirit) afforded 2 (2.80 g, 79%)
1
as a slightly yellow oil. H NMR (400 MHz, CDCl ) δ 8.56 (ddd,
3
J = 4.9, 1.8, 0.9 Hz, 1H), 8.26 (br s, 1H), 7.60 (ddd, J = 8.0, 7.4, 1.8
Hz, 1H), 7.47 (dt, J = 8.1, 1.0 Hz, 1H), 7.14 (ddd, J = 7.4, 4.9, 1.1
Hz, 1H), 4.09 (s, 2H), 3.67ꢀ3.56 (m, 2H), 2.97ꢀ2.87 (m, 2H).
13
C NMR (101 MHz, CDCl ) δ 166.3, 159.1, 150.2, 137.0,
3
2
2
its pendants or, more simply, by turning the acetate groups of
121.5, 121.4, 42.9, 38.9, 37.6. LRMS (ESI) m/z 265 (40%,
23
37 + 37 +
DOTA into amides with chiral amines.
[M[ Cl]H] ), 263 (100, [M[ Cl]H] ).
0
00
In the following, we present a new cyclen tag with three chiral
amide groups and one nonchiral pendant for attachment to a
cysteine residue of the target protein. We show that the three chiral
amide pendants are sufficient to maintain a single enantiomeric
conformation, as evidenced by a single set of peaks in the protein
NMR spectrum, and that, even at 45 °C, the tag is free of
complications arising from slow conformational exchange phe-
2,2 ,2 -(10-(2-Oxo-2-(2-(pyridin-2-yldisulfanyl)ethylamino)-
ethyl)-1,4,7,10-tetraazacyclododecane-1,4,7-triyl)tris(N-((S)-
1-phenylethyl)acetamide) (C1). A mixture containing 3 (1.40 g,
2.1 mmol), 2 (1.00 g, 3.8 mmol), and DIPEA (610 μL, 3.8 mmol)
in acetonitrile (50 mL) was stirred at room temperature for 48 h.
The resultant white precipitate was filtered and dried to afford C1
(0.65 g, 35%) as a white solid. Evaporation of the filtrate and
purification of the residue by flash chromatography (90:10:1
20
nomena that have been reported for a different chiral DOTA tag.
DCM/MeOH/NH ) afforded a further crop of the product
3
1
(
1
0.78 g, 41%). H NMR (400 MHz, DMSO) δ 8.46ꢀ8.42 (m,
’
EXPERIMENTAL PROCEDURES
H), 8.28 (t, J = 5.8 Hz, 1H), 8.19 (d, J = 8.1 Hz, 1H), 8.15 (d, J =
Tag Synthesis—(S)-2-Bromo-N-(1-phenylethyl)acetamide
1). Bromoacetyl bromide (12.7 g, 63 mmol) was added dropwise
8.1 Hz, 2H), 7.83ꢀ7.76 (m, 1H), 7.73 (dt, J = 8.1, 1.0 Hz, 1H),
(
7.33ꢀ7.25 (m, 12H), 7.24ꢀ7.16 (m, 4H), 5.00ꢀ4.86 (m, 3H),
2
119
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Bioconjugate Chemistry
ARTICLE
Figure 1. Synthesis of C1 tag and lanthanide complexes. Conditions: (i) DIPEA, chloroform, RT, O/N; (ii) DIPEA, acetonitrile, RT, 2 d; (iii)
acetonitrile, reflux, O/N.
3
.41ꢀ3.32 (m, 2H), 3.07ꢀ2.96 (m, 6H), 2.93ꢀ2.83 (m, 4H),
thiol was as above for ArgN followed by passage over a PD10
column equilibrated with degassed NMR buffer (50 mM HEPES
pH 7.5). A 15-fold excess of tag was added from a 20 mM stock
solution in water to an approximately 50 μM solution of the
protein and left for 3 h at room temperature and then overnight
at 3 °C. Excess tag was removed by passage over a PD10 column
and the sample concentrated using a Millipore ultrafilter to a final
protein concentration of about 70 μM.
13
2
.75ꢀ2.42 (m, 16H), 1.43ꢀ1.28 (m, J = 8.8, 7.1 Hz, 9H).
C
NMR (101 MHz, DMSO) δ 170.6, 169.4, 169.4, 159.1, 149.6,
1
5
44.4, 144.3, 137.7, 128.2, 126.6, 126.0, 126.0, 121.2, 119.3, 58.1,
7.9, 57.6, 53.6, 53.3, 52.9, 47.6, 37.6, 37.4, 22.2, 22.1. HRMS
+
(ESI): m/z: calcd for[M+H] C H N O S : 882.4517, found:
47 64 9 4 2
8
82.4555.
Formation of Metal Complexes. As binding of lanthanides to
DOTA-type cyclen derivatives is extremely slow and requires
heating, we did not attempt to exchange the metal ion from
tagged protein samples. To produce protein samples with
different lanthanides, the C1 tag was prepared with different
NMR Spectra. NMR spectra were recorded at 25 °C on 600
MHz NMR spectrometers equipped with cryogenic probes
(Bruker Avance and Varian INOVA in the case of ArgN and
15
ubiquitin(A28C), respectively). N-HSQC, 3D HNCA, and
HN(CO)CA spectra were recorded from differently tagged
3+
3+
3+ 3+
paramagnetic and diamagnetic ions (Tb , Tm , Yb , Y ) by
heating the tag in acetonitrile with a molar equivalent of the
respective metal triflate salts overnight at 80 °C, followed by
freezeꢀdrying to afford off-white powders. Stock solutions of the
C1ꢀlanthanide complexes were prepared in water with concen-
trations of up to 20 mM.
15
ArgN. N-fast-HSQC spectra were recorded from differently
15
tagged ubiquitin(A28C), and a 3D NOESYꢀ N-HSQC spec-
3
+
trum was recorded from the ubiquitin(A28C)ꢀC1-Y complex.
Resonance assignments of paramagnetic NMR spectra were
25
supported by the program Numbat to fit Δχ tensors to a set of
unambiguously assigned PCSs and predict the PCSs of un-
assigned paramagnetic cross-peaks in several rounds of Δχ
tensor fitting and resonance assignments.
15
13
Tagging Reaction. Uniformly N/ C-labeled samples of
the N-terminal domain of the E. coli arginine repressor (ArgN)
24
were produced on M9 minimum medium as described.
.2 mM solution of ArgN in NMR buffer (20 mM MES, pH 6.6)
A
0
Fitting of Δχ Tensors. Δχ tensors were fitted to the first
conformer of the NMR structure of ArgN (PDB accession code
was reduced by 5 equiv of DTT and subsequently washed with
NMR buffer using a Millipore ultrafilter with a molecular weight
cutoff of 3 kDa. The solution was added to a 3-fold excess of tag in
NMR buffer to yield a total volume of 0.5 mL and the mixture was
incubated at room temperature for 5 h. The reaction mixture was
washed again with NMR buffer and concentrated to a final
volume of 0.5 mL.
24
26,27
1AOY ) following a previously described protocol.
Briefly,
the structure of ArgN with the C1 tag was modeled using
the enantiomeric mirror image of the crystal coordinates of the
3+
DOTA-tetraamide Gd complex with 1-phenylethyl amine
28
(DOTAMPh, CSD accession code EQOZUF ). One of the
1-phenylethyl groups of the DOTAMPh molecule was changed
to a 2-thioethyl group, which was connected via a disulfide bond
1
5
Uniformly N-labeled samples of the mutant A28C of human
15
6
ubiquitin were prepared as described. DTT reduction of the
to the sulfur of Cys68 of ArgN. 4 ꢁ10 different conformations
2
120
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Bioconjugate Chemistry
ARTICLE
were generated by randomly changing the χ and χ angles of
1
2
Cys68 and the dihedral angles of the disulfide bond and ethylene
tether, using dihedral angles of 60°, ꢀ60°, and 180° for the CꢀC
bonds and dihedral angles of ꢀ90° or 90° for the SꢀS bond, with
an uncertainty range of (10°. Conformations producing steric
clashes with the protein were excluded. The remaining 2287
conformers were used to fit Δχ tensors to the experimental
PCSs. The final Δχ tensor was taken to be the one obtained with
the conformation producing the best least-squares fit to the
experimentally observed PCS data. As a measure of the precision
of the Δχ-tensors and metal positions, the fits to each of the 2287
model conformers were repeated 100 times, omitting 20% of the
25
data from each metal ion in a Monte Carlo protocol.
Fitting of Δχ tensors to the ubiquitin mutant A28C followed
the same protocol as for ArgN. The fits used the first conformer
29
of the NMR structure (PDB accession code 1D3Z ). 2897 con-
formers were obtained that were free of steric clashes between tag
and protein.
1
5
Figure 2. Superimposition of N-HSQC spectra of 0.2 mM solutions
1
3
15
’
RESULTS
of uniformly C/ N-labeled N-terminal domain of the E. coli arginine
3+
repressor ligated at Cys68 with the C1 tag loaded with Y (black peaks),
3
+
3+
Synthesis of the C1 Tag and Its Lanthanide Complexes.
Figure 1 shows the synthetic route used to prepare the “con-
jugation ready” C1 tag from 1,4,7,10-tetraazacyclododecane
Tb (red), or Tm (blue). All spectra were recorded at 25 °C on a
600 MHz NMR spectrometer in a 20 mM MES buffer, pH 6.6. Selected
diamagnetic cross-peaks are labeled with their resonance assignments
and connected by lines with their paramagnetic partners.
(cyclen). This involved the sequential attachment of the three
chiral amide groups and one nonchiral pendant via reaction with
α-haloacetamide derivatives of (S)-1-phenylethanamine and 2-
displaced the cross-peaks along almost straight lines. Supporting
Information Table S2 shows the experimentally determined
PCSs of ubiquitin(A28C).
(
pyridin-2-yldisulfanyl)ethanamine, respectively. Although chro-
matography was required at both stages to remove impurities, the
synthesis avoided the need for protection groups and afforded
the tag in good overall yield. The lanthanide complexes of C1
were readily prepared by refluxing 1:1 mixtures of the ligand
and metal triflate salts, Ln(OTf) (Ln = Dy , Tb , Tm , Yb ,
Y ), in acetonitrile overnight.
PCS Measurements and Δχ Tensors. Pronounced PCSs
were observed in the complexes of ArgN and ubiquitin(A28C)
3+
3+
3+
3+
tagged with C1 containing Dy , Tb , Tm , or Yb . All
samples produced high-quality NMR spectra (Figure 2 and
Supporting Information Figure S2). There was no evidence for
conformational heterogeneity of the tag or slow conformational
3+
3+
3+
3+
3
3
+
Tagging Reaction and Protein NMR Assignments. The C1
tag was attached to ArgN by adding the protein to a 3-fold excess
of tag rather than adding tag to protein, to avoid the possibility of
disulfide-bond formation between different protein molecules
which could arise from disulfide exchange. The reaction repro-
ducibly delivered tagged ArgN with greater than 95% yield.
Samples were produced with tags loaded with different lantha-
3+
exchange, neither in the C1ꢀYb tag alone (Figure S1) nor in
3+
the ubiquitin(A28C) mutant tagged with C1ꢀYb and mea-
sured with high signal-to-noise ratio (Figure S3). No minor
conformational species appeared at 45 °C (Figures S1 and S3).
Δχ tensors were fitted to the experimental PCSs using the first
24
conformer of the NMR structure of ArgN with the covalent
structure of the tags crafted onto it via a disulfide bond linkage to
Cys68. Table 1 shows the parameters of the fitted Δχ tensors. The
tensors were sufficiently large to generate PCSs of up to 0.13 ppm at
the site most remote from Cys68 (the amide proton of Asp39 in the
C1(Tb)-tagged protein which is 32 Å from the site of the metal ion).
3
+
3+
3+
3+
3+
nides (Dy , Tb , Tm , Yb ) or a diamagnetic metal (Y ).
The C1 tag was attached to the ubiquitin mutant A28C using a
similar protocol or a large excess of tag as described in the
Experimental Procedures. Reaction yields increased with the
duration of incubation (from hours to days) with the C1 tag. An
overnight reaction at 3 °C (i.e., conditions suitable for less stable
proteins) yielded 80ꢀ90% of tagged ubiquitin. Even the use of a
large excess of tag produced no evidence for disulfide mediated
protein dimerization.
2
9
Fits of the Δχ tensors to the NMR structure of ubiquitin
produced smaller tensors than in the case of ArgN (Supporting
Information Table S3), suggesting that the exposure of Cys28 on
the surface of the protein permits a greater amplitude of motion
for the tag (Figure S4; the calculated solvent accessibility of
Cys68 of ArgN is about 7% and is 22% for Cys28 of ubiquitin-
(A28C)). As in the case of ArgN, however, the C1 tag produced
significantly larger Δχ tensors than a previously used NTA
The backbone NMR resonances of paramagnetic and diamag-
netic ArgN-C1 complexes were assigned using 3D HNCA and
HN(CO)CA spectra. The resonance assignments obtained are
compiled in Supporting Information Table S1. Chemical shift
changes in ArgN and ubiquitin(A28C) tagged with diamagnetic
α
α
derivative (N ,N -bis(carboxymethyl)-L-cysteine) that is a much
3
+
15
Y
tags were limited to amides in the vicinity of the cysteine
smaller molecule with a shorter linker.
residue, indicating that the tags did not significantly alter the
Back-calculated and experimental PCSs correlated closely for
ArgN and ubiquitin (Figure 3). Remarkably, the correlation was
better for ubiquitin than for ArgN despite the greater conforma-
tional freedom of the C1 tag on the surface of ubiquitin,
indicating that tag mobility is a lesser impediment to good fits
of PCS data than structure quality.
protein structures. The resonance assignments of the diamag-
3+
netic ubiquitin(A28C)ꢀC1ꢀY complex were established by a
1
5
3
D NOESYꢀ N-HSQC spectrum. For either protein, reso-
nance assignments of the paramagnetic samples were aided
by the fact that the PCSs produced by different lanthanides
2
121
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Bioconjugate Chemistry
ARTICLE
a
Table 1. Δχ Tensors of Different Metal Ions Bound to ArgN-C1
ꢀ
32
3
ꢀ32
3
b
metal ion
Δχax /10
m
Δχrh /10
m
Q
tensor axis
coordinates of tensor axes
3
+
Dy
ꢀ29
ꢀ11
0.08
0.14
0.21
0.18
x
y
z
x
y
z
x
y
z
x
y
z
ꢀ0.302
0.941
ꢀ0.248
0.077
0.921
0.329
0.210
ꢀ0.965
0.261
0.001
0.322
0.880
0.350
0.388
0.855
0.344
(
(
ꢀ42/ꢀ25)
(ꢀ13/ꢀ7)
ꢀ0.152
ꢀ0.235
ꢀ0.935
ꢀ0.249
ꢀ0.833
0.439
0.966
3
+
Tb
ꢀ27
ꢀ37/ꢀ21)
ꢀ4
(ꢀ9/ꢀ1)
ꢀ0.064
ꢀ0.240
0.969
3
+
Tm
37
12
ꢀ0.450
ꢀ0.183
0.874
(
(
26/59)
(3/24)
ꢀ0.337
ꢀ0.808
0.495
3
+
Yb
13
3
ꢀ0.444
ꢀ0.154
0.883
10/19)
(0/7)
ꢀ0.320
a
25
24
The tensors are listed in their unique tensor representation (UTR) as obtained by fitting of the PCSs of Table S1 to ArgN (PDB ID 1AOY model 1)
3+
3+
3+
3+
simultaneously using the PCSs induced by Dy , Tb , Tm , and Yb and a common metal position. The fits used PCSs of amide protons in
structurally well-defined regions of the protein (residues 8ꢀ20, 26ꢀ37, 43ꢀ52). The covalent structure of the tag was taken into account as described in
the main text. The orientations of the tensor axes are given as unit vectors with respect to the origin (0, 0, 0). Uncertainty ranges (shown in brackets) were
obtained by a Monte Carlo error analysis that randomly omitted 20% of the PCSs (100 trials for each of 2287 modeled tag conformations). The
coordinates of the common metal position in the best fit using 100% of the PCSs were (13.705, 11.580, 0.445). A small fraction (9%) of the best solutions
in the Monte Carlo trials positioned the metal in a group about 4 Å from this position. These solutions were omitted from the error analysis. In the
b
remaining solutions, the metal was displaced by <1.9 Å. Q-factor calculated as root-mean-square deviation between measured and predicted PCSs
divided by the root-mean-square of the measured PCSs. The Q-factor of the simultaneous four-metal fit is 0.16.
1
’
DISCUSSION
800 MHz NMR spectrometer, with D
RDCs of up to
HN
3+
2
0.3 Hz measured for ArgN-C1(Tm ) at 25 °C. Maximal
Our results indicate that the C1 tag reliably produces PCSs
1
D
RDC values greater than 20 Hz are hallmarks of the best
HN
following attachment to single cysteine residues. This is an
important advantage over DPA tags
tags
8,20,31,32
1
3,14,27
lanthanide tags currently available.
Under the conditions
1
or IDA and NTA
1
5,16
of our RDC measurements, a maximal D
0 Hz would be predicted for the Δχ value of 37 ꢁ 10
value of about
HN
32 3
that often generate vanishingly small PCSs if the metal
ꢀ
2
m
ax
ion is not coordinated by an additional carboxyl group from the
protein, as mobility of the tether to the protein invariably reduces
the magnitude of the PCSs observed in the protein. The
reliability with which the C1 tag generates PCSs can be attributed
to its bulkiness, which limits the amplitude of reorientational
motions of the DOTA moiety relative to the protein. For com-
parison, no PCSs could be generated with lanthanides in the
ubiquitin (A28C) mutant tagged with the much smaller 4MDPA
tag (data not shown), despite there being only a single rotatable
bond in the linker between the lanthanide and the disulfide bond.
Reliable generation of PCSs in the absence of structural informa-
tion is an essential prerequisite before the power of PCS data can
be harnessed for 3D structure determinations of proteins.
Generation of sizable PCSs in the target protein is greatly
aided by an intrinsically large Δχ tensor of the lanthanide tag.
Remarkably large paramagnetic shifts have been reported for the
3+
associated with ArgN-C1(Tm ) (Table 1). This close agree-
ment between predicted and experimental maximal RDC values
would suggest that the tag has little conformational freedom.
Large uncertainty ranges, however, were associated with the Δχ
tensor determinations, which may be attributed to the fairly long
distance of the metal from the protein surface (8 Å from the
sulfur of the cysteine residue; Figure 4) and similar directions of
the principal tensor axes of the different metals (Table 1).
In contrast to the situation encountered with RDCs, where the
average of different alignment tensors can be represented by a
single average tensor, an ensemble of different Δχ tensors from
different metal positions cannot be approximated well by a single
average tensor, at least not close to the metal ion, since PCSs are
distance-dependent. At greater distances from the paramagnetic
center, however, the approximation by an average Δχ tensor is
much better, as different tag conformations have a smaller
relative influence on the distance between nuclear spins and
paramagnetic center. Therefore, even if mobility of the tag leads
to a reduced average Δχ tensor, this tensor still yields useful
structural restraints at greater distances from the metal ion, as
long as it fits the experimentally observed PCSs in those regions
of the protein. The large intrinsic Δχ tensor associated with the
C1 tag thus makes it a useful tool for protein structure analysis
even in the presence of substantial averaging as observed for the
ubiquitin A28C mutant. Notably, PCSs could be measured for
almost all peaks of ubiquitin (Supporting Information Figure S2)
and even using all PCSs from residues 2ꢀ72 produced a better fit
of back-calculated versus experimental PCSs for ubiquitin than
for ArgN for which only a NMR structure determined by NOEs
and J couplings is available (Figure 3).
2
7
3
+
Yb complex of the symmetrical parent compound of the C1
tag, DOTAMPh, which has 1-phenylethylacetamide pendants
23
attached to all four nitrogens of the cyclen ring. As the cyclen
protons are separated from the lanthanide by only a few bonds, it
is unclear, however, which fraction of the paramagnetic shifts in
DOTAMPh must be attributed to contact shifts rather than
PCSs. In a protein tagged with C1, however, contact shifts can be
neglected for protons that are sufficiently far from the metal to
yield observable NMR signals. Remarkably, the Δχ value of 13 ꢁ
ax
ꢀ
32
3
3+
1
0
m determined for the ArgN-C1(Yb ) complex (Table 1)
is almost two times greater than any that has been reported for
Yb complexes of calbindin D , CLaNP-5.2, lanthanide
binding peptides, or DPA tags.
residual dipolar couplings (RDC) could be observed on an
3
+
30
8,9
9k
1
9
13,19,27
Correspondingly large
2
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Bioconjugate Chemistry
ARTICLE
unrealistically large distance of the metal from the protein can
often be quite readily compensated for by a correspondingly
larger tensor. To avoid such artifacts, we used a fitting procedure
that explicitly restricts metal coordinates to positions that are
compatible with the covalent structure of the tag and its steric
requirements. Unless the tag is completely immobilized, how-
ever, the particular tag conformation producing the best fit must
not be taken too literally, as the fitting does not consider the
different rotamers of the linker moiety that may lead to the
average metal position found.
The C1 tag carries the same ethylene thio linker as the
20
previously published cyclen tags DOTA-M8 and CLaNP-
8
5
.1. In the case of the DOTA-M8 tag, 15ꢀ20% of the tag was
observed to assume a second conformation at 25 °C, which was
tentatively attributed to cisꢀtrans isomerization of the amide
2
0
bond of the tether. This percentage rose to 50% at 50 °C. In
contrast, no second conformational species was reported for the
8
CLaNP-5.1 tag and we found no evidence for slow conforma-
tional exchange in the C1 tag either, not even at 45 °C
(
Supporting Information Figures S1 and S3). The C1 tag is thus
suitable for measurements at elevated temperatures.
For the CLaNP-5.1 tag, data have been reported only for a
3+
8
single metal ion (Yb ) and protein site, making comparisons
difficult. We would expect the C1 tag to be less prone to PCS
averaging because of its greater bulkiness. Compared to the even
18,19
bulkier lanthanide binding peptides,
the C1 tag, like most
DOTA-type tags, binds lanthanide ions much more tightly, is
more resistant to unfolding under nonphysiological conditions,
and is easier to use, as it is synthesized with an activated dis-
ulfide bond. The high lanthanide affinity of DOTA-type tags
(
dissociation constants of 100 pM have been reported for
lanthanide complexes of cyclen with N-acetamide pendants,
33
DOTAM ) is a decisive advantage for experiments with metal-
loproteins, where a kinetically less inert metal binding tag could
lead to equilibration between the metals of the protein and the
tag, for studies of compounds that display high natural affinities
for lanthanides, such as DNA and RNA, and for EPR experiments
34
Figure 3. Correlations between back-calculated and experimental
pseudocontact shifts of (A) the N-terminal domain of the E. coli arginine
repressor and (B) ubiquitin obtained by the fits of Table 1 and
Supporting Information Table S3, respectively.
3+
3+
with Gd that depend on the absence of unbound Gd ions.
The parent compound of the C1 tag, DOTAMPh, forms a
23
square antiprismatic complex that completely encases the
lanthanide ions except for a single coordination site for a water
35
molecule. The poor accessibility of the lanthanide is an
advantage in interaction studies, where more solvent-exposed
metal ions could produce misleading results by direct interac-
tions between the metal and the interaction partner. The C1 tag
thus carries great promise for lead compound development by
fragment-based drug design, where PCSs observed for weakly
binding fragments can be used to define their binding site,
36,37
orientation, and structure.
Figure 4. Model of the N-terminal domain of the E. coli arginine
repressor with a C1-lanthanide complex bound to Cys68. The model
represents the best fit of the PCSs to the first conformer of the NMR
Using different lanthanides in the same protein-tag construct
generates Δχ tensors of different magnitude, but the orientations
13,30,38
24
of the principal axes of the tensors usually vary little.
It is a
structures (PDB ID 1AOY ). The structure of the tag was derived from
3+
the crystal coordinates of a DOTA-tetraamide Gd complex as
described in the main text. The lanthanide ion is represented by a
magenta sphere. The disulfide bond of the linker is highlighted in yellow.
The N- and C-termini are labeled. The side chain of Cys68 and the tag
are shown in a heavy atom representation.
distinct possibility that a tag produced with opposite chirality (using
(R)-1-phenylethanamine instead of (S)-1-phenylethanamine) may
generate different tensor orientations relative to the protein. PCS
data measured with both enantiomeric tags could thus deliver
two independent sets of restraints to position nuclear spins
relative to the protein.
Δχ tensor fits easily generate large uncertainty ranges for
metal tags on protein surfaces, as the metal position must be
determined along with five other tensor parameters (Euler
angles and axial and rhombic components). For example, an
In conclusion, the new lanthanide tag presented here displays
an attractive combination of properties, including reliable gen-
eration of PCSs, conformational homogeneity, and tight lantha-
nide binding. We anticipate that it will become an important tool
2
123
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Bioconjugate Chemistry
ARTICLE
for assessing the structure of biological macromolecules by NMR
spectroscopy.
Engineering encodable lanthanide-binding tags into loop regions of
proteins. J. Am. Chem. Soc. 133, 808–819.
(13) Man, B., Su, X.-C., Liang, H., Simonsen, S., Huber, T., Messerle,
B. A., and Otting, G. (2010) 3-Mercapto-2,6-pyridinedicarboxylic acid, a
small lanthanide-binding tag for protein studies by NMR spectroscopy.
Chem.—Eur. J. 16, 3827–3832.
’
ASSOCIATED CONTENT
Supporting Information. H NMR spectrum of the
1
S
b
(14) Su, X.-C., Man, B., Beeren, S., Liang, H., Simonsen, S., Schmitz,
3
+
C1ꢀYb complex, chemical shifts of ArgN tagged with C1
C., Huber, T., Messerle, B. A., and Otting, G. (2008) A dipicolinic acid
tag for rigid lanthanide tagging of proteins and paramagnetic NMR
spectroscopy. J. Am. Chem. Soc. 130, 10486–10487.
15
loaded with different metal ions, N-HSQC spectra of ubiquitin
A28C) tagged with C1 loaded with different metal ions, PCSs of
(
ubiquitin (A28C) tagged with C1 metal complexes, Δχ tensors
of ubiquitin (A28C) tagged with C1 metal complexes, model of
the C1 tag bound to Cys28 in ubiquitin (A28C). This material is
available free of charge via the Internet at http://pubs.acs.org.
(15) Swarbrick, J. D., Ung, P., Su, X.-C., Maleckis, A., Chhabra, S.,
Huber, T., Otting, G., and Graham, B. (2011) Engineering of a bis-
chelator motif into a protein α-helix for rigid lanthanide binding and
paramagnetic NMR spectroscopy. Chem. Commun. 47, 7368–7370.
(16) Swarbrick, J. D., Ung, P., Chhabra, S., and Graham, B. (2011)
An iminodiacetic acid based lanthanide binding tag for paramagnetic
exchange NMR spectroscopy. Angew. Chem., Int. Ed. 50, 4403–4406.
’
AUTHOR INFORMATION
(
17) Lu, Y., Berry, S. M., and Pfister, T. D. (2001) Engineering novel
Corresponding Author
metalloproteins: design of metal-binding sites into native protein
scaffolds. Chem. Rev. 101, 3047–3080.
*
Gottfried Otting, Research School of Chemistry, Australian
National University, Canberra, ACT 0200 (Australia). Fax:
61-2-61250750. E-mail: gottfried.otting@anu.edu.au.
(18) Su, X.-C., Huber, T., Dixon, N. E., and Otting, G. (2006) Site-
+
specific labelling of proteins with a lanthanide-binding tag. ChemBio-
Chem 7, 1469–1474.
(
19) Su, X.-C., McAndrew, K., Huber, T., and Otting, G. (2008)
’
ACKNOWLEDGMENT
Lanthanide-binding peptides for NMR measurements of residual dipolar
couplings and paramagnetic effects from multiple angles. J. Am. Chem.
Soc. 130, 1681–1687.
Financial support by the Australian Research Council, includ-
ing a Future Fellowship to T. H., is gratefully acknowledged.
(20) H €a ussinger, D., Huang, J., and Grzesiek, S. (2009) DOTA-M8:
an extremely rigid, high-affinity lanthanide chelating tag for PCS NMR
spectroscopy. J. Am. Chem. Soc. 131, 14761–14767.
’
REFERENCES
(
1) Pintacuda, G., John, M., Su, X.-C., and Otting, G. (2007) NMR
(21) Parker, D., Dickins, R. S., Puschmann, H., Crossland, C., and
Howard, J. A. K. (2002) Being excited by lanthanide coordination
complexes: aqua species, chirality, excited-state chemistry, and exchange
dynamics. Chem. Rev. 102, 1977–2010.
(22) Woods, M., Botta, M., Avedano, S., Wang, J., and Sherry, A. D.
(2005) towards the rational design of MRI contrast agents: a practical
approach to the synthesis of gadolinium complexes that exhibit optimal
water exchange. Dalton Trans. 2005, 3829–3837.
(23) Dickins, R. S., Parker, D., Bruce, J. I., and Tozer, D. J. (2003)
Correlation of optical and NMR spectral information with coordination
variation for axially symmetric macrocyclic Eu(III) and Yb(III) com-
plexes: axial donor polarisability determines ligand field and cation
donor preference. Dalton Trans. 2003, 1264–1271.
structure determination of protein-ligand complexes by lanthanide
labelling. Acc. Chem. Res. 40, 206–212.
(
2) Otting, G. (2008) Prospects for lanthanides in structural biology
by NMR. J. Biomol. NMR 42, 1–9.
3) Bertini, I., Luchinat, C., and Parigi, G. (2002) Magnetic suscept-
ibility in paramagnetic NMR. Prog. Nucl. Magn. Reson. Spectrosc. 40,
49–273.
4) Keizers, P. H. J., Mersinli, B., Reinle, W., Donauer, J., Hiruma, Y.,
(
2
(
Hannemann, F., Overhand, M., Bernhardt, R., and Ubbink, M. (2010) A
solution model of the complex formed by adrenodoxin and adrenodoxin
reductase determined by paramagnetic NMR spectroscopy. Biochemistry
4
9, 6846–6855.
(
5) Saio, T., Yokochi, M., Kumeta, H., and Inagaki, F. (2010) PCS-
(24) Sunnerhagen, M., Nilges, M., Otting, G., and Carey, J. (1997)
Solution structure of the DNA-binding domain and model for the
binding of multifunctional arginine repressor to DNA. Nat. Struct. Biol.
4, 819–826.
based structure determination of protein-protein complexes. J. Biomol.
NMR 46, 271–280.
(
6) Keizers, P. H. J., and Ubbink, M. (2011) Paramagnetic tagging
for protein structure and dynamics analysis. Prog. Nucl. Magn. Reson.
Spectrosc. 58, 88–96.
(25) Schmitz, C., Stanton-Cook, M. J., Su, X.-C., Otting, G., and
Huber, T. (2008) Numbat: an interactive software tool for fitting
Δχ-tensors to molecular coordinates using pseudocontact shifts. J. Biomol.
NMR 41, 179–189.
(
7) Otting, G. (2010) Protein NMR using paramagnetic ions. Annu.
Rev. Biophys. 39, 387–405.
8) Keizers, P. H. J., Desreux, J. F., Overhand, M., and Ubbink, M.
2007) Increased paramagnetic effect of a lanthanide protein probe by
two-point attachment. J. Am. Chem. Soc. 129, 9292–9293.
9) Keizers, P. H. J., Saragliadis, A., Hiruma, Y., Overhand, M., and
(
(26) Nguyen, T. H. D., Ozawa, K., Stanton-Cook, M., Barrow, R.,
(
Huber, T., and Otting, G. (2011) Pseudocontact shifts in protein NMR
2
+
spectra generated using a genetically encoded Co -binding amino acid.
(
Angew. Chem., Int. Ed. 50, 692–694.
(27) Jia, X., Maleckis, A., Huber, T., and Otting, G. (2011) 4,4 -
0
Ubbink, M. (2008) Design, synthesis, and evaluation of a lanthanide
chelating protein probe: CLaNP-5 yields predictable paramagnetic
effects independent of environment. J. Am. Chem. Soc. 130, 14802–
Dithiobis-dipicolinic acid: a small and convenient lanthanide-binding tag
for protein NMR spectroscopy. Chem.—Eur. J. 17, 6830–6836.
(28) Parker, D., Puschmann, H., Batsanov, A. S., and Senanayake, K.
(2003) Structural analysis of nine-coordinate lanthanide complexes:
steric control of the metal-water distance across the series. Inorg. Chem.
42, 8646–8651.
(29) Cornilescu, G., Marquardt, J. L., Ottiger, M., and Bax, A. (1998)
Validation of protein structure from anisotropic carbonyl chemical shifts
in a dilute liquid crystalline phase. J. Am. Chem. Soc. 120, 6836–6837.
(30) Bertini, I., Janik, M. B. L., Lee, Y. M., Luchinat, C., and Rosato,
A. (2001) Magnetic susceptibility tensor anisotropies for a lanthanide
ion series in a fixed protein matrix. J. Am. Chem. Soc. 123, 4181–4188.
14812.
(
10) Vlasie, M. D., Comuzzi, C., van den Nieuwendijk, A. M.,
Prud ^e ncio, M., Overhand, M., and Ubbink, M. (2007) Long-range-
distance NMR effects in a protein labeled with a lanthanide-DOTA
chelate. Chem.—Eur. J. 13, 1715–1723.
(11) Saio, T., Yokochi, M., and Inagaki, F. (2009) Two-point
anchoring of a lanthanide-binding peptide to a target protein enhances
the paramagnetic anisotropic effect. J. Biomol. NMR 44, 157–166.
(12) Barthelmes, K., Reynolds, A. M., Peisach, E., Jonker, H. R. A.,
DeNunzio, N. J., Allen, K. N., Imperiali, B., and Schwalbe, H. (2011)
2
124
dx.doi.org/10.1021/bc200353c |Bioconjugate Chem. 2011, 22, 2118–2125

Bioconjugate Chemistry
ARTICLE
(
31) Su, X.-C., and Otting, G. (2010) Paramagnetic labelling of
proteins and oligonucleotides for NMR. J. Biomol. NMR 46, 101–112.
32) Su, X.-C., and Otting, G. (2011) Erratum to: Paramagnetic
labelling of proteins and oligonucleotides for NMR. J. Biomol. NMR
0, 99–100.
33) Maumela, H., Hancock, R. D., Carlton, L., Reibenspies, J. H.,
(
5
(
and Wainwright, K. P. (1995) The amide oxygen as a donor group. Metal
ion complexing properties of tetra-N-acetamide substituted cyclen: a
crystallographic, NMR, molecular mechanics, and thermodynamic
study. J. Am. Chem. Soc. 117, 6698–6707.
(34) Potapov, A., Yagi, H., Huber, T., Jergic, S., Dixon, N. E., Otting,
G., and Goldfarb, D. (2010) Nanometer scale distance measurements in
3+
proteins using Gd spin labeling. J. Am. Chem. Soc. 132, 9040–9048.
35) Aime, S., Barge, A., Batsanov, A. S., Botta, M., Castelli, D. D.,
(
Fedeli, F., Mortillaro, A., Parker, D., and Puschmann, H. (2002)
Controlling the variation of axial water exchange rates in macrocyclic
lanthanide(III) complexes. Chem. Commun. 2002, 1120–1121.
(
36) John, M., Pintacuda, G., Park, A. Y., Dixon, N. E., and Otting, G.
2006) Structure determination of protein-ligand complexes by trans-
ferred paramagnetic shifts. J. Am. Chem. Soc. 128, 12910–12916.
37) Zhuang, T. D., Lee, H. S., Imperiali, B., and Prestegard, J. H.
2008) Structure determination of a Galectin-3-carbohydrate complex
using paramagnetism-based NMR constraints. Protein Sci. 17, 1220–1231.
38) Pintacuda, G., Park, A. Y., Keniry, M. A., Dixon, N. E., and
(
(
(
(
Otting, G. (2006) Lanthanide labeling offers fast NMR approach to 3D
structure determinations of protein-protein complexes. J. Am. Chem. Soc.
128, 3696–3702.
2
125
dx.doi.org/10.1021/bc200353c |Bioconjugate Chem. 2011, 22, 2118–2125