Thiol-ene reaction: A versatile tool in site-specific labelling of proteins with chemically inert tags for paramagnetic NMR

Article abstract of DOI:10.1039/c2cc17900h

Site-specific tagging of proteins with paramagnetic lanthanides generates valuable long-range structure restraints for structural biology by NMR spectroscopy. We show that the thiol-ene addition reaction offers a powerful tool for tagging proteins in a chemically stable manner with very small lanthanide tags. The Royal Society of Chemistry 2012.

Full text of DOI:10.1039/c2cc17900h

View Article Online / Journal Homepage / Table of Contents for this issue
Dynamic Article Links
ChemComm
Cite this: Chem. Commun., 2012, 48, 2704–2706
www.rsc.org/chemcomm
COMMUNICATION
Thiol–ene reaction: a versatile tool in site-specific labelling of proteins
with chemically inert tags for paramagnetic NMRw
Qing-Feng Li,a Yin Yang,a Ansis Maleckis,b Gottfried Otting*b and Xun-Cheng Su*ab
Received 18th December 2011, Accepted 19th January 2012
DOI: 10.1039/c2cc17900h
Site-specific tagging of proteins with paramagnetic lanthanides
generates valuable long-range structure restraints for structural
biology by NMR spectroscopy. We show that the thiol–ene
addition reaction offers a powerful tool for tagging proteins in
a chemically stable manner with very small lanthanide tags.
Paramagnetic lanthanide ions provide an extraordinarily valuable
tool for structural studies of biomolecules by NMR spectroscopy,1
including structure determinations of protein–protein and protein–
Fig. 1 Lanthanide binding tags of DPA derivates containing a vinyl
ligand complexes,2 and the study of conformational exchange.3 As
or maleimide group.
proteins rarely bind lanthanides specifically, generation of
paramagnetic data usually depends on site-specific tagging
with lanthanide-chelating reagents.4 Pseudocontact shifts
and the ligation product is stable in the presence of DTT or TCEP.
Significant PCSs were observed with lanthanides in two proteins,
the N-terminal domain of the E. coli arginine repressor (ArgN)9
and human ubiquitin.
(PCSs) require immobilization of the lanthanides with respect
to the protein, leading to many recent efforts to improve the
rigidity of the paramagnetic tags.5 At present, chemically
We synthesized two lanthanide tags, L1 and L2 (Fig. 1, details
synthesized tags are almost invariably attached to the protein
of synthesis of L1 and L2 are provided in the ESIw). L1 is a vinyl
derivative of dipicolinic acid (DPA). L2 is a maleimide derivative.
Both are Michael receptors owing to a double bond that is
activated by an electron-withdrawing group. L1 reacts specifically
with the thiol group of cysteine (see ESIw for model reactions) and
no reaction was observed for the amino group of amino acids or
the side chain of lysine at pH 7.8. Iodoacetamide and maleimide
derivatives of nitroxide radicals are widely used in EPR studies,10
but corresponding lanthanide tags are not as useful for NMR
studies as iodoacetamide tags can react with side chain amino
groups of lysine and the reaction of maleimide with cysteine
generates a new chiral center which leads to diastereomeric
protein–tag conjugates. Indeed, two species became apparent in
the 15N-HSQC spectrum of ArgN–L2 after addition of Tm3+
(Fig. S2, ESIw). Therefore, we subsequently focused on L1.
Quantitative ligation of ArgN with L1 was achieved by
incubating the protein with an eight-fold excess of L1 at room
temperature and pH 7.6 for 24 hours. Low-molecular weight
compounds were subsequently separated from the ligated product
by ion-exchange chromatography (yield 4 80%). The resulting
ligation product formally resembles the previously described
ArgN–4MMDPA complex,5a except that one of the sulfur atoms
is replaced by a methylene group. Titration of ArgN–L1 with
paramagnetic lanthanide ions produced large chemical shift
changes in the 15N-HSQC spectrum (Fig. 2) similar in direction
and magnitude to those obtained with ArgN–4MMDPA.5a
Different from ArgN–L2, only a single paramagnetic species
by formation of one or two disulfide bonds. This precludes the
subsequent use of reducing agents or high pH, and makes
these constructs incompatible with thiol-containing ligands.
They have also been observed to suffer during long storage.6
While the insertion of a lanthanide binding peptide into the
loop of a protein would avoid this complication,5e there are
only a few suitable fusion sites in a protein if any. Here we
present the first chemically synthesized tag that attaches to
cysteine residues spontaneously and by formation of an inert
thioether by virtue of the thiol–ene reaction (Scheme 1) that is
well established in polymer science7 and has so far been rarely
applied in protein chemistry.8 The tagging reaction is thiol specific
Scheme 1 Site-specific tagging proteins via thiol–ene reaction in
aqueous solution.
a State-Key Laboratory of Elemento-organic Chemistry,
Nankai University, Tianjin, 300071, China.
E-mail: xunchengsu@nankai.edu.cn
b Research School of Chemistry, Australian National University,
Canberra, ACT 0200, Australia. E-mail: go@rsc.anu.edu.au
w Electronic supplementary information (ESI) available. See DOI:
10.1039/c2cc17900h
c
2704 Chem. Commun., 2012, 48, 2704–2706
This journal is The Royal Society of Chemistry 2012

View Article Online
Fig. 2 Superimposition of 15N-HSQC spectra of uniformly 15N-labeled
ArgN derivatized with L1 at Cys68 in the presence of a 1 : 1 mixture of
Y3+ and Yb3+ (black) and in the presence of Y3+ alone (red). The ratio
of lanthanide to protein was about 1.2 : 1. The spectra were recorded at
25 1C and pH 6.5 at a 1H NMR frequency of 600 MHz.
Fig. 3 Superimposition of 15N-HSQC spectra of a 0.08 mM solution
of uniformly 15N-labeled ubiquitin T22C derivatized with L1 in the
presence of a 1 : 1 mixture of Y3+ and Yb3+ (red) and in the presence
of Y3+ (black). The ratio of lanthanide to protein was about 1.2 : 1.
The spectra were recorded at 25 1C and pH 6.4 at a 1H NMR
frequency of 600 MHz.
Table 1 Dw tensor parameters of ArgN–L1 and ubiquitin–L1 complexed
with Dy3+, Tb3+, Tm3+, and Yb3+, respectivelya
Dy3+ Tb3+
Tm3+
Yb3+
ArgN–L1
Dwax 23.7
Dwrh 11.6
17.3 (12.9) 11.0 (12.2) 5.7 (5.7)
7.1 (6.3)
19.2
8.4
5.6 (6.9)
13.8
7.5
2.6 (3.8)
7.7
2.2
Ubiquitin T22C–L1 Dwax
Dwrh
a
The tensor parameters are in units of 10À32 m3. The tensor para-
meters determined for the corresponding lanthanide complexes of
ArgN–4MMDPA5a are shown in brackets for comparison.
was observed as the molar ratio of lanthanide to protein was
increased from 0.2 to 1.5. Four sets of PCSs were measured for
Tb3+, Dy3+, Tm3+ and Yb3+, and the respective Dw tensor
parameters were determined using the program Numbat11
with the first conformer of the NMR structure of ArgN9
(Table 1). The metal position was found at a distance of
4.3 A to one of the oxygens of the E21 side chain, which is
1.6 A longer than in ArgN–4MMDPA.5a In view of the
flexibility of the poorly determined side chain conformation
of E21, coordination to E21 appears as the likely mechanism
that restricts the mobility of the metal ion. The magnitudes of
the Dw tensors observed in ArgN–L1 are similar to those
obtained with ArgN–4MMDPA.
Fig. 4 Ribbon drawing of ubiquitin showing the metal position
(black sphere) determined by fitting the PCSs of the ubiquitin
T22C–L1-lanthanide complexes to the ubiquitin structure (PDB ID:
1UBQ), and the respective distances to the side chains of D21, E24,
and the g-carbon of T22.
We subsequently tested a different attachment site in a different
protein by constructing the T22C mutation in ubiquitin. T22 is
located at the beginning of the a-helix and has a solvent exposed
side chain. Two acidic residues, D21 and E24, have side chains
within 7 A of T22. Incubation of ubiquitin T22C with L1
produced ubiquitin T22C–L1 in high yield (485%). Large
PCSs of up to 0.5 ppm were observed even in the presence of
the weakly paramagnetic lanthanide Yb3+ (Fig. 3). Although
protein concentrations below 0.1 mM had to be used to avoid
intermolecular aggregation in the presence of lanthanides,
the complexes yielded high-quality 15N-HSQC spectra. PCSs
were measured for Tb3+, Tm3+ and Yb3+, and the respective
Dw tensor parameters were determined using the crystal structure
of ubiquitin (pdb code: 1UBQ).12 The magnitudes of the
Dw tensors of the ubiquitin T22C–L1 lanthanide complexes
indicated that the tag is similarly immobilized as in ArgN.
The calculated metal position was 7.1 A from the Cg atom of
T22 and 4.7 A from one of the carboxyl oxygens of D21,
implying additional coordination to D21 (Fig. 4). This example
shows that the L1 tag is sufficiently small to allow immobiliza-
tion of the lanthanide ion between two neighboring residues
in the amino acid sequence.
Incubation of ArgN–L1 and ubiquitin T22C–L1 with five
equivalents of DTT or TCEP produced no changes in the
15N-HSQC spectra of these proteins irrespective of the
presence or absence of lanthanide metal ions. In addition,
the solution of ubiquitin T22C–L1 with lanthanide was stable
at room temperature and pH 8.2 over five days without
significant chemical shift changes, confirming the chemical
stability of the thioethers formed in the thiol–ene addition
reaction. L1 reacts with cysteine residues more slowly than
previously published thiol-activated lanthanide binding tags.4b
This can be advantageous in the presence of buried cysteine
residues. For example, no reaction was observed with the buried
c
This journal is The Royal Society of Chemistry 2012
Chem. Commun., 2012, 48, 2704–2706 2705

View Article Online
cysteines of E. coli peptidylprolyl isomerase B with a 50-fold
excess of L1 at room temperature and pH 7.6 for three days.
In conclusion, the new L1 tag presents a powerful and
convenient way of tagging proteins with paramagnetic lanthanides
free of chirality issues, generating a stable ligation product.
Although requiring prior knowledge of the structure of the
protein, the possibility to immobilize the lanthanide between
two neighboring residues by a very small and inert tag makes it a
most attractive tool for structural biology by NMR spectroscopy.
This work was supported by National Science Foundation
of China (grant agreement number 21073101 to X. C. S.) and
the Australian Research Council (to G. O.).
37, 79–95; (d) M. A. Hass, P. H. Keizers, A. Blok, Y. Hiruma and
M. Ubbink, J. Am. Chem. Soc., 2010, 132, 9952–9953.
4 (a) F. Rodriguez-Castaneda, P. Haberz, A. Leonov and C. Griesinger,
Magn. Reson. Chem., 2006, 44, S10–S16; (b) X. C. Su and G. Otting,
J. Biomol. NMR, 2009, 46, 101–112; (c) P. H. Keizers and M. Ubbink,
Prog. Nucl. Magn. Reson. Spectrosc., 2011, 58, 88–96.
5 (a) X. C. Su, B. Ma, S. Beeren, H. Liang, S. Simonsen, C. Schmitz,
T. Huber, B. Messerle and G. Otting, J. Am. Chem. Soc., 2008,
130, 10486–10487; (b) P. H. Keizers, A. Saragliadas, Y. Hiruma,
M. Overhand and M. Ubbink, J. Am. Chem. Soc., 2008, 130,
14802–14812; (c) D. Haussinger, J. R. Huang and S. Grzesiek,
¨
J. Am. Chem. Soc., 2009, 131, 14761–14767; (d) T. Saio, K. Ogura,
M. Yokochi, Y. Kobashigawa and F. Inagaki, J. Biomol. NMR,
2009, 44, 157–166; (e) K. Barthelmes, A. M. Reynolds, E. Peisach,
H. R. Jonker, N. J. DeNunzio, K. N. Allen, B. Imperiali and
H. Schwalbe, J. Am. Chem. Soc., 2011, 133, 808–819;
(f) J. D. Swarbrick, P. Ung, X. C. Su, A. Maleckis, S. Chhabra,
T. Huber, G. Otting and B. Graham, Chem. Commun., 2011, 47,
7368–7370; (g) F. Peters, M. Maestre-Martinez, A. Leonov, L.
Kovacic, S. Beckers, R. Boelens and C. Griesinger, J. Biomol. NMR,
2011, 51, 329–337.
6 J. Iwahara, C. Tang and G. M. Clore, J. Magn. Reson., 2007, 184,
185–195.
7 (a) K. L. Killops, L. M. Campos and C. J. Hawer, J. Am. Chem. Soc.,
2008, 130, 5062–5064; (b) C. E. Hoye, A. B. Lowe and C. N. Bowan,
Chem. Soc. Rev., 2010, 39, 1355–1387.
8 (a) J. E. T. Corrie, J. Chem. Soc., Perkin Trans. 1, 1994, 2975–2982;
(b) V. Hong, A. A. Kislukhin and M. G. Finn, J. Am. Chem. Soc.,
2009, 131, 9986–9994; (c) E. M. Sletten and C. R. Bertozzi,
J. Am. Chem. Soc., 2009, 48, 6974–6998.
9 M. Sunnerhagen, M. Nilges, G. Otting and J. Carey, Nat. Struct. Biol.,
1997, 4, 819–826.
10 P. A. Kosen, Methods Enzymol., 1989, 177, 86–120.
11 C. Schmitz, M. J. Stanton-Cook, X. C. Su, G. Otting and
T. Huber, J. Biomol. NMR, 2008, 41, 179–189.
12 S. Vijay-Kumar, C. E. Bugg and W. J. Cook, J. Mol. Biol., 1987,
194, 531–544.
Notes and references
1 (a) I. Bertini, C. Luchinat and G. Parigi, Prog. NMR Spectrosc.,
2002, 40, 249–273; (b) G. Pintacuda, M. John, X. C. Su and
G. Otting, Acc. Chem. Res., 2007, 40, 206–212; (c) G. Otting,
J. Biomol. NMR, 2008, 42, 1–9.
2 (a) M. John, G. Pintacuda, A. Y. Park, N. E. Dixon and G. Otting,
J. Am. Chem. Soc., 2006, 128, 12910–12916; (b) G. Pintacuda,
A. Y. Park, M. A. Keniry, N. E. Dixon and G. Otting, J. Am.
Chem. Soc., 2006, 128, 3696–3702; (c) E. Balogh, D. Wu, G. Zhou
and M. Gochin, J. Am. Chem. Soc., 2009, 131, 2821–2823;
(d) P. H. Keizers, B. Mersinli, W. Reinle, J. Donauer, Y. Hiruma,
F. Hannemann, M. Overhand, R. Bernhardt and M. Ubbink,
Biochemistry, 2010, 49, 6846–6855; (e) T. Saio, K. Ogura,
M. Yokochi and F. Inagaki, J. Biomol. NMR, 2010, 46, 271–280.
3 (a) I. Bertini, C. Del Bianco, I. Gelis, N. Katsaros, C. Luchinat,
G. Parigi, M. Peana, A. Provenzani and M. A. Zoroddu,
Proc. Natl. Acad. Sci. U. S. A., 2004, 101, 6841–6846; (b) X.
Wang, S. Srisailam, A. Yee, A. Lemak, C. Arrowsmith, J. H.
Prestegard and F. Tian, J. Biomol. NMR, 2007, 39, 53–61;
(c) C. Eichmuller and N. R. Skrynnikov, J. Biomol. NMR, 2006,
¨
c
2706 Chem. Commun., 2012, 48, 2704–2706
This journal is The Royal Society of Chemistry 2012