2′-Alkynylnucleotides: A Sequence- and Spin Label-Flexible Strategy for EPR Spectroscopy in DNA

Article abstract of DOI:10.1021/jacs.6b05421

Electron paramagnetic resonance (EPR) spectroscopy is a powerful method to elucidate molecular structure through the measurement of distances between conformationally well-defined spin labels. Here we report a sequence-flexible approach to the synthesis of double spin-labeled DNA duplexes, where 2′-alkynylnucleosides are incorporated at terminal and internal positions on complementary strands. Post-DNA synthesis copper-catalyzed azide-alkyne cycloaddition (CuAAC) reactions with a variety of spin labels enable the use of double electron-electron resonance experiments to measure a number of distances on the duplex, affording a high level of detailed structural information.

Full text of DOI:10.1021/jacs.6b05421

Subscriber access provided by UNIV OF CALIFORNIA SAN DIEGO LIBRARIES
Communication
2#-Alkynylnucleotides: A sequence- and spin label-
flexible strategy for EPR spectroscopy in DNA
Marius M. Haugland, Afaf H. El-Sagheer, Rachel J. Porter, Javier
Pena, Tom Brown, Edward A. Anderson, and Janet E. Lovett
J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/jacs.6b05421 • Publication Date (Web): 13 Jul 2016
Downloaded from http://pubs.acs.org on July 16, 2016
Just Accepted
“Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted
online prior to technical editing, formatting for publication and author proofing. The American Chemical
Society provides “Just Accepted” as a free service to the research community to expedite the
dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts
appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been
fully peer reviewed, but should not be considered the official version of record. They are accessible to all
readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered
to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published
in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just
Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor
changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers
and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors
or consequences arising from the use of information contained in these “Just Accepted” manuscripts.
Journal of the American Chemical Society is published by the American Chemical
Society. 1155 Sixteenth Street N.W., Washington, DC 20036
Published by American Chemical Society. Copyright © American Chemical Society.
However, no copyright claim is made to original U.S. Government works, or works
produced by employees of any Commonwealth realm Crown government in the course
of their duties.

Page 1 of 5
Journal of the American Chemical Society
1
2
3
4
5
6
7
8
9
2′-Alkynylnucleotides: A sequence- and spin label-flexible
strategy for EPR spectroscopy in DNA
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Marius M. Haugland,† Afaf H. El-Sagheer,†¶ Rachel J. Porter,† Javier Peña,‡ Tom Brown,*,† Ed-
ward A. Anderson*,† and Janet E. Lovett*§
† Department of Chemistry, Chemistry Research Laboratory, University of Oxford, Oxford, OX1 3TA, U.K.
‡ Departamento de Química Orgánica, Universidad de Salamanca, Plaza de los Caídos 1–5, 37008 Salamanca, Spain
¶ Chemistry Branch, Faculty of Petroleum and Mining Engineering, Suez University, Suez 43721, Egypt
§ SUPA, School of Physics and Astronomy, University of St Andrews, North Haugh, St Andrews, KY16 9SS, U.K.
Supporting Information Placeholder
through the incorporation of 2′-alkynylnucleosides into DNA
(2). As well as tolerating standard DNA synthesis conditions,
this sequence-flexible approach enables post-DNA synthesis
CuAAC click reactions to install a variety of spin labels.
DEER experiments on double spin-labeled duplexes afford an
array of distance measurements, which
ABSTRACT: EPR spectroscopy is a powerful method to elu-
cidate molecular structure through the measurement of dis-
tances between conformationally well-defined spin labels.
Here we report a sequence-flexible approach to the synthesis
of double spin-labeled DNA duplexes, where 2′-
alkynylnucleosides are incorporated at terminal and internal
positions on complementary strands. Post-DNA synthesis
copper-catalyzed azide-alkyne cycloaddition (CuAAC) reac-
tions with a variety of spin labels enables the use of double
electron-electron resonance experiments to measure a num-
ber of distances on the duplex, affording a high level of de-
tailed structural information.
Previous work:
This work:
O
O
Base
modification
Ribose
modification
• baseꢀflexible
siteꢀdirected spin
labeling (SDSL)
N
NH
O
N
O
1
Base
O
O
NH
N
O
O
O
P
X
spin
linker
O
label
O
O
O
O
P
N
O
O
O
O
N
N
• label flexibility
• postꢀDNA synthesis
labeling strategy
H
O
P
HN
N
N
spin
label
spin
label
O
O
2
Ç
S
The development of selective and accurate methods for
visualizing biomolecules is essential for understanding the
relationship between structure and activity in biological sys-
tems. Electron paramagnetic resonance (EPR) spectroscopy
is rapidly emerging as a powerful tool in this context.1 The
pulsed EPR technique double electron-electron resonance
(DEER) is particularly useful for applications to biopolymers,
where the dipolar coupling between two paramagnetic cen-
ters enables the measurement of interspin distances of 15 to
>100 Å, as well as spin label orientations.2
Figure 1. Examples of SDSL strategies: Cytidine analogue Ç,
2 -linked spin labels (1), and the 2 -ethynyl-2 -
deoxynucleotide SDSL approach (2).
generate a detailed structural 'map' of the duplex, and reveal
the importance of spin label structure on distance distribu-
tion.
The synthesis of 2′-ethynyl-2′-deoxyuridine (3, Scheme 1)
built on the elegant silicon-tethered radical alkyne transfer
chemistry developed by Shuto et al.7 Starting with cyclouri-
dine 4, protected 2′-iodonucleoside 5 was obtained in two
steps. Mild activation of aminosilane 6 with iodomethane
greatly enhanced yields in the subsequent challenging silyla-
tion of the 3′-hydroxyl group of 5.8 The resulting silyl ether 7
was immediately submitted to the radical atom transfer step,
where the reported conditions7 (0.3 equiv. BEt3, toluene, Ar)
led to incomplete conversion and side reactions. However,
the reaction proceeded smoothly when performed in ben-
zene with trace amounts of air, which delivered 2′-
alkynyluridine derivative 3 in high yield on multigram scale.
3 was also converted to the 2′-alkynylcytidine derivative 8.9
In biopolymers without natural paramagnetic centers (e.g.
metal ions), spin labels must be introduced in a site-specific
fashion. These should be conformationally restricted to max-
imize accuracy in EPR experiments, but equally should not
perturb the structure or function of the biopolymer. A variety
of strategies have been developed for the site-directed spin
labeling (SDSL) of nucleic acids,3 including modification of
the nucleobase,2a,4 phosphodiester,5 and ribose framework.6
For example, the cytidine analog Ç (Figure 1) gives rigid oli-
gonucleotide structures and detailed angular information,4e,4f
but inherently restricts sequence generality. 2′-(Thio)urea6c-f
and propargyl ether-derived triazole6b linkers (1) achieve
ribose labeling; despite greater sequence flexibility, the for-
mer requires hydrolytically-sensitive labeling reagents (e.g.
iso(thio)cyanates), whilst the latter are relatively mobile
which reduces the accuracy of distance measurements. Here
we describe a new approach to oligonucleotide spin-labeling
The preparation of 2′-ethynyl-2′-deoxyadenosine 9 proved
more challenging. This began with arabinoadenosine 10,
which was converted to 2′-iodonucleoside 11 using a disilox-
ane protection strategy. After N-benzoylation, desilylation,
and tritylation of the 5′-hydroxyl, equivalent chemistry used
ACS Paragon Plus Environment

Journal of the American Chemical Society
Page 2 of 5
in the synthesis of 3 was deployed. The alkynylsilyl ether
amidites 13 for incorporation into internal positions of DNA
1
2
3
derived from 12 was less stable than uridine analog 7, and
was submitted to the radical atom transfer without chroma-
tographic purification, which gave 9 in reasonable yield.
oligonucleotides (Scheme 2). Additionally, 3 was resin-
mounted via succinylation of the 3′-alcohol (14), thereby en-
abling
modification
of
the
3′-
4
5
The 2′-alkynylnucleoside derivatives 3, 8 and 9 were con-
verted to the corresponding 3′-O-(cyanoethyl)phosphor-
6
7
Scheme 1. Synthesis of 2 -ethynyl-2 -deoxynucleoside derivatives.
a) TBSCl, imid., DMF
b) POCl3, NEt3,
1,2,4ꢀtriazole, MeCN;
then NH3, dioxane
O
8
9
6
NHBz
N
O
O
O
BEt3,
2,6ꢀlutidine,
benzene;
Et2N Si
MeI;
TMS
N
a) NaI, pꢀTsOH,
acetone, 50 °C
NH
NH
NH
O
O
N
HO
DMTrO
DMTrO
DMTrO
DMTrO
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
N
O
N
O
N
O
N
O
then 5, imid.,
O
O
O
O
c) BzCl, pyridine
d) TBAF, THF
b) DMTrCl, pyridine
88% (2 steps)
then TBAF
98%
DMAP, CH2Cl2
96%
OH
OH
I
O
I
OH
OH
70% (4 steps)
Si
4
5
7
3
8
TMS
6
NH2
NH2
N
NHBz
NHBz
Et2N Si
TMS
N
N
N
N
N
N
N
N
N
a) BzCl, pyridine;
then NH3
a) MeI;
HO
O
DMTrO
DMTrO
a) TIPDSꢀCl2, imid., DMF
then 12, imid., DMAP, CH2Cl2
N
N
O
N
N
N
N
O
iꢀPr2Si
O
O
OH
b) TfCl, NEt3, DMAP, CH2Cl2
c) LiI, DMF
b) TBAF, THF
c) DMTrCl, pyridine
b) BEt3, 2,6ꢀlutidine,
benzene; then TBAF
O
OH
O
I
OH
I
OH
iꢀPr2Si
10
63% (3 steps)
11
69% (3 steps)
12
51% (2 steps)
9
Scheme 2. Synthesis of spin labeled DNA duplexes.
Duplex ꢀTm (°C)
N3
N3
N3
N3
R
R
R
Duplex 6-Me
A2:BUL –4.9
AUL:B2 –3.1
A1:B2 –3.1
A2:B2 –9.8
5-Me
6-Sp
–2.3
–0.6
–0.6
–3.7
5-Sp
–4.1
In-Me
–4.0
–1.4
–0.6
–6.4
In-Et
–3.2
–1.0
+0.7
–4.3
N3
N
O
–4.9
–3.6
N
O
N
O
N
O
DMTrO
N
O
–2.1
–1.1
–8.0
R
NiꢀPr2
Base
O
P
CN
–3.5
In-Me: R = Me
In-Et: R = Et
iꢀPr2N
O
6-Me
5-Me
6-Sp
5-Sp
3, 8, 9
–10.4
iꢀPr2N
O
Nꢀmethylimidazolium
triflate, MeCN
P
O
UL:
1:
2:
5'-ACTAGCGCGGACTCTT-3'
5'-ACTAGCGCGGACTCTU-3'
5'-ACTAGCGCGGACUCTT-3'
5'-ACTAGCGCGGACTCTT-3'
CN
20
10
0
Duplex 6-Me
A1:B3 –3.1
A1:B4 –1.7
A2:B1 –5.2
A2:B3 –9.6
A2:B4 –7.1
5-Sp
–1.8
–0.8
ꢀ
In-Et
+0.1
–0.3
ꢀ
13
Strand A
a) succinic anhydride, iꢀPr2EtN,
DMAP, CH2Cl2
b) resin, DIC, HOBT, pyridine
c) C6Cl5OH
d) piperidine, THF
e) Ac2O, Nꢀmethylimidazole,
pyridine, THF
3:
O
DNA
synthesis
UL: 5'-AAGAGTCCGCGCTAGT-3'
1: 5'-AAGAGTCCGCGCTAGU-3'
2: 5'-AAGAGTCCGCGCUAGT-3'
3: 5'-AAGAGTCCGCGCTAGT-3'
4: 5'-AAGAGTCCGCGCTAGT-3'
NH
CuI (5 equiv.),
THPTA (10 equiv.),
spin label (100 equiv.),
MeCN / H2O
DMTrO
Strand B
N
O
–8.9
–6.6
–1.2
–4.3
–4.2
ꢀ
ꢀ10
ꢀ20
O
AUL:BUL
A2:B2 5ꢀMe
O
A3:B2
ꢀ
O
200 240 280 320
Wavelength (nm)
N
H
14
O
terminus of the DNA. These building blocks were successful-
ly incorporated into two complementary single stranded
DNA oligonucleotides using automated solid-phase synthe-
sis, at both terminal and internal positions (Strands A1–3 and
B1–4, Scheme 2; unlabeled strands AUL and BUL were also
prepared). To our knowledge, this is the first time a 2′-alkyne
modification has been incorporated into DNA. The versatility
of this approach is emphasized by the incorporation of the
modified nucleotide at three adjacent (different) bases on
Strand B.
performed at higher temperatures.4b,12 Tetraalkylisoindoline
spin labels such as In-Me and In-Et are also of interest due
to their enhanced rigidity, combined with high stability to-
ward reducing environments.6f,13
These spin labels were site-selectively attached to the
modified DNA via CuAAC chemistry.14 Due to the intoler-
ance of some of these nitroxyls to sodium ascorbate, modi-
fied reaction conditions were used (copper(I) iodide / tris(3-
hydroxypropyltriazolylmethyl) amine). Duplexes with vari-
ous combinations of A and B strands were formed, and the
consequence of introducing the spin label at the 2 -position
was investigated through UV denaturation studies and CD
spectroscopy.15 We first examined the structural influence of
a single internal label (A2:BUL and AUL:B2) on the DNA
duplex melting temperature (Tm).16 Internal labeling of
Strand B was consistently found to be less destabilizing, with
a moderate (up to 4 °C) dependence on the spin label identi-
ty. Little or no change from these values was seen in double
spin-labeled duplexes with terminal and internal labels (e.g.
A1:B2). However, double-internal spin labeling had a more
significant effect, and also showed the greatest variance of Tm
with spin label (up to 6.7 °C), although there appeared to be
little correlation with the size of the label. For example,
A2:B2–6-Sp showed the lowest Tm depression; this may be
due to hydrophobic stacking interactions, possibly with the
sugar to its 3 -side. CD spectroscopy confirmed that the B-
The divergent nature of the strategy facilitates the explora-
tion of the properties of different spin labels. For this we
identified a small library of six azidonitroxides (see box,
Scheme 2), which would be incorporated into the modified
oligonucleotides post-DNA synthesis.10 These spin labels
were selected to explore the competing requirements of la-
bel/duplex rigidity and minimized structural distortion, in-
cluding labels which display improved biological stability
and/or spectroscopic properties. The well-established 6-Me
and 5-Me enable evaluation of our strategy against other
methods, with 5-Me known to be more resistant to bioreduc-
tion.11 Rotation of the methyl groups in these nitroxides pro-
vides a mechanism for relaxation of excited electron spins;
this can be avoided through the use of the rigidified
bis(spirocyclohexyl) spin labels 6-Sp and 5-Sp, where longer
phase memory times (T2m) enable DEER measurements to be
ACS Paragon Plus Environment

Page 3 of 5
Journal of the American Chemical Society
form DNA helix is conserved, even in the most destabilized
thyl substituents generally exhibit a pronounced turning
1
2
3
4
5
6
7
8
duplex (Scheme 2, CD spectra of AUL:BUL and A2:B2–5-
Me).10
point in their relaxation rates as the rotation of the alkyl
groups becomes less dominant in the relaxation process;12a in
our duplexes, this was only observed for In-Et, which could
indicate that local environment is an important factor for
this effect.10,12c The 6-Sp and 5-Sp labels were observed to
have a much longer T2m at all temperatures.12
Room temperature continuous wave (CW) EPR was car-
ried out using single-labeled duplexes.10 The A1:BUL duplexes
showed greater label mobility than A2:BUL or AUL:B2, which
is consistent with labeling of a terminal nucleotide in
A1:BUL. Interestingly, marked differences in mobility were
observed between different labels in all duplexes, with 6-Me
and 5-Me exhibiting the most rapid dynamics. The T1 relaxa-
tion time and T2m were next determined for the various la-
bels in A1:BUL.10 Labels with flanking tetramethyl or tetrae-
Pulsed EPR techniques were used to measure distances be-
tween spin labels in double-labeled duplexes. The back-
ground-corrected DEER time traces for the 6-Me-labeled
duplexes typically
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Figure 2. DEER data for the spin labeled duplexes. a) DEER time traces for the series of 6-Me labeled duplexes. A3 is 5-Sp la-
beled and A2':B2' denotes a 2 -triazolylmethyl ether tether (see text). b) Distance distributions derived from DEER using Deer-
Analysis201517 for the data series shown in a. c) Distance distributions for A1:B2 duplexes. d) Distance distributions for A2:B2
duplexes.
showed one or two frequency modulations (Figure 2a), sug-
gesting that the labels occupy a relatively narrow region of
conformational space.
(Figure 2c, black curve) which suggests three distinct con-
formations of the pairs of labels. The double-internal A2:B2
duplexes (Figure 2d) replicate this phenomenon: the most
probable distances (and conformations) clearly depend on
label type. The conformational preferences correlate with the
nature of the nitroxyl-containing ring: 5-membered rings 5-
Me and 5-Sp are positioned at ~4.2 nm separation, while 6-
membered rings 6-Me and 6-Sp are at ~3.6 nm. It is also
notable that the width of the distributions appear unrelated
to the dynamics of the labels observed in the rt CW EPR
experiments: for example, the distributions for A2:B2–5-Me
and –5-Sp overlap completely, although the dynamics are
different.18 The broadest distributions were consistently ob-
tained for In-Et, despite the slow dynamics exhibited in the
CW EPR; this may imply that this sterically encumbered spin
label associates with the duplex in a less well-defined man-
ner than other labels.
The DEER-derived distance distributions obtained for this
series of duplexes (Figure 2b) show that the average hwhh of
0.45 ± 0.13 nm is broadly independent of the influence of the
label on the Tm of the duplex, and whether the label is placed
internally or at the 3′ end of the duplex (in contrast to the
measured mobilities at rt). Pleasingly, similar distance meas-
urements were observed for the pseudo-symmetric A1:B2
(red curve) and A2:B1 (black curve) duplexes, which supports
well-defined positioning of the labels. An exception was the
double internal labeled A2:B4, which gave a broader distri-
bution; however, this illustrates the value of positioning spin
labels at will to improve accuracy in the exploration of du-
plex structure. The red dashed curve in Figure 2b depicts 6-
Me attached via a 2′-O-propargyl ether-clicked triazole in
the double internal labeled A2':B2'. The hwhh increased to
0.65 nm, reflecting the increased mobility associated with
this more flexible linker, compared to 0.29 nm for A2:B2–6-
Me.6b Similar results were obtained using 5-Sp and In-Et
labels with this linker.10 For duplexes with interspin distanc-
es at which DEER is less accurate (<1.5 nm),2e line-
broadening of the frozen CW EPR spectra was used to ap-
proximate distances using 5-Sp as the spin label; this gave
values for A3:B2, A3:B3 and A3:B4 (0.9, 1.0 and 1.3 nm respec-
tively).
A representation of the B-form helical DNA of the unla-
beled duplex AUL:BUL is illustrated in Figure 3. The most
probable distances from the DEER experiments and the ap-
proximate distances from CW EPR are shown, with calculat-
ed distances between the 2 -hydroxyls in a B-form mixed
DNA/RNA model duplex depicted for comparison.19 Good
correlation is found between these modeled distances and
the measured interspin distances, particularly for internal
labels (mean deviation –0.01 ± 0.06 nm for DEER measure-
ments). Notably, the measured A3:B4 distance (1.30 nm) is
significantly greater than A3:B2 or A3:B3 (~1.0 nm), despite
similar calculated 2 -OH–2 -OH distances. This may imply
that the spin labels are oriented toward the 3 terminus in a
directionally well-defined manner, rather than projecting
into solution.6d,6f The duplexes labeled at the 3 termini con-
sistently afford distances greater than the crude model
(mean deviation 0.49 ± 0.31 nm for 6-Me labeled duplexes).
The DEER distance measurement, and hence label envi-
ronment, was found to depend on the structure of the spin
label. For example, measurements on the various A1:B2 du-
plexes (Figure 2c) afforded similar distances, with 5-Sp giv-
ing the narrowest distribution. Interestingly, the distance
distribution for In-Me showed three well-resolved maxima
ACS Paragon Plus Environment

Journal of the American Chemical Society
Page 4 of 5
(2) (a) Halbmair, K.; Seikowski, J.; Tkach, I.; Höbartner, C.; Sezer,
The hwhh of these distributions suggests that the labels
nonetheless adopt well-defined conformations.
D.; Bennati, M. Chem. Sci. 2016, 7, 3172; (b) Endeward, B.; Marko, A.;
Denysenkov, V. P.; Sigurdsson, S. T.; Prisner, T. F. Methods Enzymol.
2015, 564, 403; (c) El Mkami, H.; Norman, D. G. Methods Enzymol.
2015, 564, 125; (d) Borbat, P. P.; Freed, J. H. Struct. Bond. 2013, 152, 1;
(e) Banham, J. E.; Baker, C. M.; Ceola, S.; Day, I. J.; Grant, G. H.;
Groenen, E. J. J.; Rodgers, C. T.; Jeschke, G.; Timmel, C. R. J. Magn.
Reson. 2008, 191, 202; (f) Lovett, J. E.; Bowen, A. M.; Timmel, C. R.;
Jones, M. W.; Dilworth, J. R.; Caprotti, D.; Bell, S. G.; Wong, L. L.;
Harmer, J. Phys. Chem. Chem. Phys. 2009, 11, 6840.
1
2
3
4
5
6
7
8
9
(3) Shelke, S. A.; Sigurdsson, S. T. Nucleic Acids Mol. Biol. 2016, 31,
159.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
(4) (a) Duss, O.; Yulikov, M.; Jeschke, G.; Allain, F. H. T. Nat.
Commun. 2014, 5, 3669; (b) Babaylova, E. S.; Ivanov, A. V.; Malygin,
A. A.; Vorobjeva, M. A.; Venyaminova, A. G.; Polienko, Y. F.;
Kirilyuk, I. A.; Krumkacheva, O. A.; Fedin, M. V.; Karpova, G. G.;
Bagryanskaya, E. G. Org. Biomol. Chem. 2014, 12, 3129; (c) Ding, P.;
Wunnicke, D.; Steinhoff, H. J.; Seela, F. Chem. Eur. J. 2010, 16, 14385;
(d) Sicoli, G.; Wachowius, F.; Bennati, M.; Höbartner, C. Angew.
Chem. Int. Ed. 2010, 49, 6443; (e) Schiemann, O.; Cekan, P.; Margraf,
D.; Prisner, T. F.; Sigurdsson, S. T. Angew. Chem. Int. Ed. 2009, 48,
3292; (f) Barhate, N.; Cekan, P.; Massey, A. P.; Sigurdsson, S. T.
Angew. Chem. Int. Ed. 2007, 46, 2655; (g) Schiemann, O.; Piton, N.;
Plackmeyer, J.; Bode, B. E.; Prisner, T. F.; Engels, J. W. Nat. Protoc.
2007, 2, 904.
(5) (a) Popova, A. M.; Kálai, T.; Hideg, K.; Qin, P. Z. Biochemistry
2009, 48, 8540; (b) Cai, Q.; Kusnetzow, A. K.; Hubbell, W. L.;
Haworth, I. S.; Gacho, G. P. C.; Van Eps, N.; Hideg, K.; Chambers, E.
J.; Qin, P. Z.; Price, E. a. Nucleic Acids Res. 2006, 34, 4772; (c)
Nguyen, P. H.; Popova, A. M.; Hideg, K.; Qin, P. Z. BMC Biophys.
2015, 8:6.
(6) (a) Shevelev, G. Y.; Krumkacheva, O. A.; Lomzov, A. A.;
Kuzhelev, A. A.; Rogozhnikova, O. Y.; Trukhin, D. V.; Troitskaya, T.
I.; Tormyshev, V. M.; Fedin, M. V.; Pyshnyi, D. V.; Bagryanskaya, E.
G. J. Am. Chem. Soc. 2014, 136, 9874; (b) Flaender, M.; Sicoli, G.;
Fontecave, T.; Mathis, G.; Saint-Pierre, C.; Boulard, Y.; Gambarelli,
S.; Gasparutto, D. Nucleic Acids Symp. Ser. 2008, 52, 147; (c)
Edwards, T. E.; Okonogi, T. M.; Robinson, B. H.; Sigurdsson, S. T. J.
Am. Chem. Soc. 2001, 123, 1527; (d) Ward, R.; Keeble, D. J.; El Mkami,
H.; Norman, D. G. ChemBioChem 2007, 8, 1957; (e) Kim, N. K.;
Bowman, M. K.; DeRose, V. J. J. Am. Chem. Soc. 2010, 132, 8882; (f)
Saha, S.; Jagtap, A. P.; Sigurdsson, S. T. Chem. Commun. 2015, 51,
13142.
Figure 3. A representation of the B-form helical DNA (Maes-
tro 10.5, Schrödinger, LLC). The A-strand backbone is shown
in green and the B-strand in blue. Labeled positions are rep-
resented in orange. Modal distances from DEER distance
measurements (see Figure 2) and powder CW EPR spectra
(denoted *) are shown. Distances in brackets were measured
between the 2 -hydroxyl O atoms of the model (Pymol 1.4,
Schrödinger, LLC).
In conclusion, we have developed a sequence-flexible
method for oligonucleotide spin labeling. 2 -Alkynyl-
nucleosides were incorporated into a 16-mer DNA duplex at
various positions using standard solid phase synthesis, and
functionalized with a range of azide-containing nitroxides.
Pulsed and CW EPR experiments were used to determine
dynamic and electron spin relaxation properties of the labels,
and to measure interspin distances, with the structure of the
radical probe influencing the positioning of the spin label on
the duplex. The structural information obtained demon-
strates the versatility of this approach in exploring the struc-
ture of nucleic acids, including applications beyond SDSL.
ASSOCIATED CONTENT
Supporting Information
(7) (a) Sukeda, M.; Ichikawa, S.; Matsuda, A.; Shuto, S. Angew.
Chem. Int. Ed. 2002, 41, 4748; (b) Sukeda, M.; Ichikawa, S.; Matsuda,
A.; Shuto, S. J. Org. Chem. 2003, 68, 3465.
(8) Ohshita, J.; Iwata, A.; Kanetani, F.; Kunai, A.; Yamamoto, Y.;
Matui, C. J. Org. Chem. 1999, 64, 8024.
(9) Divakar, K. J.; Reese, C. B. J. Chem. Soc. Perkin Trans. 1 1982,
1171.
(10) See the Supporting Information for details.
Supporting Information: Experimental conditions, copies of
spectroscopic data, characterization of oligonucleotides.
Corresponding Authors
tom.brown@chem.ox.ac.uk;
ed-
ward.anderson@chem.ox.ac.uk; jel20@st-andrews.ac.uk
(11) Couet, W. R. R.; Brasch, R. C. C.; Sosnovsky, C.; Lukszo, J.;
Prakash, I.; Gnewech, C. T. T.; Tozer, T. N. N. Tetrahedron 1985, 41,
1165.
Notes
The authors declare no competing financial interests.
(12) (a) Kathirvelu, V.; Smith, C.; Parks, C.; Mannan, M. A.; Miura,
Y.; Takeshita, K.; Eaton, S. S.; Eaton, G. R. Chem. Commun. 2009,
454; (b) Kirilyuk, I. A.; Polienko, Y. F.; Krumkacheva, O. A.;
Strizhakov, R. K.; Gatilov, Y. V.; Grigor'ev, I. A.; Bagryanskaya, E. G.
J. Org. Chem. 2012, 77, 8016; (c) Kuzhelev, A. A.; Strizhakov, R. K.;
Krumkacheva, O. A.; Polienko, Y. F.; Morozov, D. A.; Shevelev, G. Y.;
Pyshnyi, D. V.; Kirilyuk, I. A.; Fedin, M. V.; Bagryanskaya, E. G. J.
Magn. Reson. 2016, 266, 1.
(13) (a) Jagtap, A. P.; Krstic, I.; Kunjir, N. C.; Hänsel, R.; Prisner, T.
F.; Sigurdsson, S. T. Free Radical Res. 2015, 49, 78; (b) Paletta, J. T.;
Pink, M.; Foley, B.; Rajca, S.; Rajca, A. Org. Lett. 2012, 14, 5322.
(14) (a) El-Sagheer, A. H.; Brown, T. Chem. Soc. Rev. 2010, 39,
1388; (b) Amblard, F.; Cho, J. H.; Schinazi, R. F. Chem. Rev. 2009,
109, 4207.
ACKNOWLEDGMENT
TB and AHE-S thank the BBSRC (sLOLA grant
BB/J001694/1). EAA thanks the EPSRC (EP/K005391/1,
EP/M019195/1). JEL thanks the EPSRC (EP/LO22044/1), and
the Royal Society for a University Research Fellowship and
grant RF120645. We thank the Wellcome Trust for the Q-
band EPR spectrometer (099149/Z/12/Z), the EPSRC National
EPR Service for spectrometer time, and Dr Hassane El Mka-
mi for technical assistance.
REFERENCES
(15) (a) 2′-Triazoles derived from 2′-azidonucleotides are well-
tolerated in nucleic acids. See: Fauster, K.; Hartl, M.; Santner, T.;
Aigner, M.; Kreutz, C.; Bister, K.; Ennifar, E.; Micura, R. ACS Chem.
(1) (a) Klare, J. P.; Steinhoff, H. J. Photosynth. Res. 2009, 102, 377;
(b) Ward, R.; Schiemann, O. Struct. Bond. 2013, 152, 249.
ACS Paragon Plus Environment

Page 5 of 5
Journal of the American Chemical Society
Biol. 2012, 7, 581; (b) Sau, S. P.; Hrdlicka, P. J. J. Org. Chem. 2012, 77,
5.
(16) The duplex AUL:BUL displayed a Tm value of 68.1 °C. The
typical error (standard deviation) in Tm measurement is ±0.3-0.5 °C.
See the Supporting Information.
(17) Jeschke, G.; Chechik, V.; Ionita, P.; Godt, A.; Zimmermann,
H.; Banham, J.; Timmel, C. R.; Hilger, D.; Jung, H. Appl. Magn.
Reson. 2006, 30, 473.
(18) The differences between label types were further compared
through labeling the A1, A2 and B2, B3, B4 duplexes with 6-Me, 5-Sp
and In-Et; see the Supporting Information.
(19) More accurate interpretation of the positioning and
orientation of the different spin labels through modeling and / or X-
ray crystallographic studies will be reported in due course.
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Table of contents image:
ACS Paragon Plus Environment