Unlocked nucleic acids with a pyrene-modified uracil: Synthesis, hybridization studies, fluorescent properties and i-motif stability

Article abstract of DOI:10.1002/cbic.201300567

The synthesis of two new phosphoramidite building blocks for the incorporation of 5-(pyren-1-yl)uracilyl unlocked nucleic acid (UNA) monomers into oligonucleotides has been developed. Monomers containing a pyrene-modified nucleobase component were found to destabilize an i-motif structure at pH 5.2, both under molecular crowding and noncrowding conditions. The presence of the pyrene-modified UNA monomers in DNA strands led to decreases in the thermal stabilities of DNA/DNA and DNA/RNA duplexes, but these duplexes' thermal stabilities were better than those of duplexes containing unmodified UNA monomers. Pyrene-modified UNA monomers incorporated in bulges were able to stabilize DNA/DNA duplexes due to intercalation of the pyrene moiety into the duplexes. Steady-state fluorescence emission studies of oligonucleotides containing pyrene-modified UNA monomers revealed decreases in fluorescence intensities upon hybridization to DNA or RNA. Efficient quenching of fluorescence of pyrene-modified UNA monomers was observed after formation of i-motif structures at pH 5.2. The stabilizing/destabilizing effect of pyrene-modified nucleic acids might be useful for designing antisense oligonucleotides and hybridization probes. Copyright

Full text of DOI:10.1002/cbic.201300567

CHEMBIOCHEM
FULL PAPERS
DOI: 10.1002/cbic.201300567
Unlocked Nucleic Acids with a Pyrene-Modified Uracil:
Synthesis, Hybridization Studies, Fluorescent Properties
and i-Motif Stability
Pavla Perlꢀkovꢁ,[a, b] Kasper K. Karlsen,[a] Erik B. Pedersen,[a] and Jesper Wengel*[a]
The synthesis of two new phosphoramidite building blocks for
the incorporation of 5-(pyren-1-yl)uracilyl unlocked nucleic acid
(UNA) monomers into oligonucleotides has been developed.
Monomers containing a pyrene-modified nucleobase compo-
nent were found to destabilize an i-motif structure at pH 5.2,
both under molecular crowding and noncrowding conditions.
The presence of the pyrene-modified UNA monomers in DNA
strands led to decreases in the thermal stabilities of DNA*/DNA
and DNA*/RNA duplexes, but these duplexes’ thermal stabili-
ties were better than those of duplexes containing unmodified
UNA monomers. Pyrene-modified UNA monomers incorporat-
ed in bulges were able to stabilize DNA*/DNA duplexes due to
intercalation of the pyrene moiety into the duplexes. Steady-
state fluorescence emission studies of oligonucleotides con-
taining pyrene-modified UNA monomers revealed decreases in
fluorescence intensities upon hybridization to DNA or RNA.
Efficient quenching of fluorescence of pyrene-modified UNA
monomers was observed after formation of i-motif structures
at pH 5.2. The stabilizing/destabilizing effect of pyrene-modi-
fied nucleic acids might be useful for designing antisense oli-
gonucleotides and hybridization probes.
Introduction
Nucleic acids play key roles in numerous cellular processes in
which they adopt various structures and perform various func-
tions. Chemical modifications can endow naturally occurring
nucleic acids with unique properties, and chemically modified
nucleic acids have therefore been widely explored for molecu-
lar diagnostics and therapeutic applications.
Unlocked nucleic acids (UNAs, 2’,3’-seco-RNAs) are acyclic
analogues of RNA that lack CÀC bonds between 2’- and 3’-
carbon atoms (Scheme 1).[1] Opening of the furanose ring leads
to increased flexibility in an UNA monomer. This has a negative
effect on the stabilities of nucleic acid duplexes,[2–6] but can im-
prove mismatch discrimination.[3,6] UNA monomers were also
found to modulate the stabilities of i-motif[7] and G-quadru-
plex[8] structures. UNA monomers therefore meet the require-
ments for applications in antisense oligonucleotides,[9,10] siRNA
constructs[11] and aptamer development.[12] Chemical synthesis
of UNA phosphoramidite building blocks is simple,[2,3] and an
UNA monomer can be incorporated into oligonucleotides
either through the O3’[2,3] or the O2’[2,4,9] atom. The sugar
moiety of an UNA monomer can easily be conjugated with, for
Scheme 1. UNA monomers.
example, ligands for complexation of metal ions[13] or attach-
ment of fluorescent labels[6] such as pyrene.
Pyrene units are widely used for oligonucleotide functionali-
zation, due to pyrene’s fluorescent properties and its aromatic
stacking and duplex intercalating abilities.[14,15]
A pyrene
moiety can be attached either to the sugar part of the nucleo-
tide or to the nucleobase. It can also substitute the nucleobase
or be used in non-nucleosidic monomers such as intercalating
nucleic acid (INA)[16] or twisted intercalating nucleic acid (TINA)
monomers.[17]
[a] Dr. P. Perlꢀkovꢁ, Dr. K. K. Karlsen, Prof. E. B. Pedersen, Prof. J. Wengel
Nucleic Acid Center, Department of Physics, Chemistry, and Pharmacy
University of Southern Denmark
Campusvej 55, 5230 Odense M (Denmark)
E-mail: jwe@sdu.dk
[b] Dr. P. Perlꢀkovꢁ
2’-Pyrene-modified UNA oligonucleotides (incorporating
monomers W or Z, Scheme 1) display increased fluorescence
upon hybridization to DNA, together with pyrene excimer
emission, and have been employed in development of molecu-
lar beacons.[18] Furthermore, introduction of the pyrene moiety
Institute of Organic Chemistry and Biochemistry
Academy of Sciences of the Czech Republic
Flemingovo nꢁm. 2, 16610 Prague 6 (Czech Republic)
Supporting information for this article is available on the WWW under
http://dx.doi.org/10.1002/cbic.201300567.
ꢂ 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemBioChem 2014, 15, 146 – 156 146

CHEMBIOCHEM
FULL PAPERS
www.chembiochem.org
into UNA monomers led to stabilization of nucleic acid duplex-
es relative to unmodified UNA, whereas the strong mismatch
discrimination capabilities of appropriately designed UNA olig-
omers were maintained.[6] The pyrene units of 2’-pyrene-modi-
fied UNA monomers were found to be located in the major
groove in the nucleic acid duplex.[6]
The positioning of a functionality into the major groove of
a nucleic acid duplex can conveniently be achieved by func-
tionalization of the 5-position of a pyrimidine nucleobase.
Direct conjugation of the pyrene with this nucleobase compo-
nent is expedient because covalent linking between the
pyrene moiety and a pyrimidine base leads to strong electron
coupling between the two aromatic units, resulting in interest-
ing fluorescence properties such as charge transfer emis-
sion.[19,20] Oligonucleotides incorporating 5-(pyren-1-yl)-2’-deox-
yuridine units have therefore been intensively used for studies
of electron-transfer processes in DNA.[21–23] Furthermore, oligo-
nucleotides containing 2’-deoxyuridine monomers with a fluo-
rescent label in the 5-position were found to change fluores-
cence properties upon formation of an i-motif structure, thus
allowing probing of i-motifs.[24]
Scheme 2. a) i: NaIO4/1,4-dioxane, H2O, RT; ii: NaBH4/1,4-dioxane, H2O, RT;
b) BzCl (Bz=benzoyl), DBU/CH2Cl2, À788C; c) pyrene-1-B(OH)2, Na2CO3, PPh3,
Pd(OAc)2/toluene/H2O, 1008C; d) P(Cl)(NiPr2)OCH2CH2CN, DIPEA/CH2Cl2, RT;
e) oligonucleotide synthesis.
Preparation of oligonucleotides (ONs) containing UNA mon-
omers bearing pyrene substituents at position 5 of the uracil
base is of interest because these monomers combine the flexi-
bility of the UNA with the stabilization effect of the pyrene
moiety in the major groove of a nucleic acid duplex. Further-
more, 5-(pyren-1-yl)uracilyl nucleotides show interesting fluo-
rescence properties.[19–23] In this report we describe the synthe-
sis of two new phosphoramidite building blocks for the in-
corporation of 5-(pyren-1-yl)uracilyl UNA monomers X and Y
(Schemes 2 and 3) into ONs. We have studied the thermal sta-
bilities and the steady-state fluorescence emission and UV/visi-
ble spectra of DNA*/DNA and DNA*/RNA duplexes of oligonu-
cleotides containing 5-pyrene-modified UNA monomers, as
well as those of DNA*/DNA and DNA*/RNA duplexes with
bulge-incorporated 5-pyrene-modified UNA monomers. The
thermal stabilities and the fluorescence and UV/visible absorp-
tion properties of i-motif-forming ONs with 5-pyrene-modified
UNA monomers are also reported.
knecht[20,23] could not be applied because of rapid hydrolysis of
the benzoyl group in the presence of NaOH, so milder condi-
tions with Na2CO3 as a base were used.[26] Even though an
excess (3 equiv) of the boronic acid was used, partial deiodina-
tion of the starting material was observed, but the pyrenyl-
substituted nucleoside 4 was obtained in 35% yield. Subse-
quent O3’-phosphitylation of nucleoside 4 by treatment with
2-cyanoethyl-N,N-diisopropylchlorophosphoramidite afforded
phosphoramidite 5 in 93% yield.
The synthesis of phosphoramidite 8 (Scheme 3) started with
silylation of the 3’-hydroxy group of nucleoside 3 by treatment
with TBSCl in pyridine, followed by removal of the 2’-O-benzoyl
group by hydrolysis with NaOH, leading to O3’-silylated nu-
cleoside 6 in 76% yield over the two steps. Subsequent
Suzuki–Miyaura cross-coupling with pyrene-1-boronic acid in
the presence of Na2CO3, PPh3 and Pd(OAc)2 in toluene/water at
1008C afforded nucleoside 7 in 57% yield. Finally, O2’-phosphi-
tylation of nucleoside 7 provided phosphoramidite 8 in 65%
yield.
Results and Discussion
Synthesis
A synthetic approach to the phosphoramidite building blocks
5 and 8 from DMT-protected 5-iodouridine 1[25] was devel-
oped. 2’,3’-Bond cleavage was accomplished by treatment with
NaIO4, and subsequent reduction of the dialdehyde intermedi-
ate with NaBH4 afforded the acyclic derivative 2 in 72% yield
(Scheme 2). Selective benzoylation with benzoyl chloride in the
presence of DBU in CH2Cl2 at À788C provided O2’-benzoate 3
in 70% yield. Formation of a by-product tentatively assigned
as the corresponding O3’-benzoate was observed, but the
compound was not isolated. A pyrene moiety was introduced
into the 5-position in benzoate 3 by means of a Suzuki–
Miyaura cross-coupling reaction with pyrene-1-boronic acid
under palladium catalysis. The conditions used by Wagen-
The phosphoramidite building blocks 5 and 8 were used in
an automated DNA synthesizer to incorporate 3’-linked mono-
mer X and 2’-linked monomer Y, respectively, into ONs [ON1–6
(Table 1) and ON7 and ON8 (Table 5, below)].
Thermal denaturation studies
The thermal stabilities (Tm values) of duplexes formed between
ONs with single or double incorporation of monomer X or Y
(DNA*) and complementary DNA or RNA were studied
(Table 1). The Tm values of DNA*/DNA and DNA*/RNA duplexes
with single incorporation of either X or Y in position 10 (ON1,
ON4) were determined to be approximately 48C lower than
ꢂ 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemBioChem 2014, 15, 146 – 156 147

CHEMBIOCHEM
FULL PAPERS
www.chembiochem.org
the presence of the pyrene moiety in the 5-position led to in-
creases of 2–118C in their thermal stabilities relative to their
counterparts containing unmodified UNA monomer (U).[6] The
stabilities of duplexes of ONs containing monomers X and Y
were found to be similar to those of duplexes containing
2’-pyrene-modified UNA monomers W and Z (Scheme 1).[6] It
follows that major groove interactions of the pyrene moiety
improve the duplex thermal stability regardless of whether the
pyrene is attached through the 5- or the 2’-position of an UNA
monomer.
Mismatches were introduced into position 11 (juxtaposition
to monomers X and Y in ON1 and ON4) in the nucleic acid du-
plexes. Tm values were determined, and DTm values relative to
fully matched duplexes of the studied ONs were calculated
(Table 1). Singly modified ONs (ON1, ON4) showed mismatch
discrimination similar to or slightly lower than that of the un-
modified duplexes. Only in the case of the DNA*/RNA duplex
of ON4—with a G:U mismatch—was the discrimination higher
than for the unmodified duplex. Mismatch discrimination capa-
bilities of modified ONs containing two incorporations of X or
Y separated by one base (ON2, ON5) were low, and the DTm
values ranged from À3 to +18C. On the other hand, when
incorporations of X and Y were nine bases apart (ON3, ON6)
mismatch discrimination was improved to 9–128C.
Mismatches were also introduced at position 10 (position di-
rectly opposite to monomers X and Y in ON1) in the duplexes
(Table 2). Poor mismatch discrimination was observed for ONs
ON1, ON2, ON4 and ON5. For these ONs the DTm values
ranged from À3 to +18C. However, the duplexes of oligonu-
cleotides ON3 and ON6, each containing two incorporations
separated by nine bases, showed highly efficient mismatch dis-
crimination (5–148C). The mismatch discrimination capabilities
of ONs containing monomers X and Y follow the same trend
as seen with other previously reported UNA monomers (U, W,
Z).[3,6]
Scheme 3. a) i: TBSCl/py, RT; ii: NaOH/MeOH, RT; b) pyrene-1-B(OH)2, Na2CO3,
PPh3, Pd(OAc)2/toluene/H2O, 1008C; c) P(Cl)(NiPr2)OCH2CH2CN, DIPEA/CH2Cl2,
RT; d) oligonucleotide synthesis.
the Tm values of the corresponding unmodified references,
showing the destabilization effect of the pyrene-modified UNA
monomers. In addition, the thermal stabilities of DNA*/DNA
and DNA*/RNA duplexes with two incorporations of X or Y
were lower than those of the unmodified duplexes. The du-
plexes in which the two insertions were separated by a single
nucleotide were less stable than those in which the two inser-
tions were nine bases apart. The Tm values of duplexes contain-
ing UNA monomers X and Y were similar to each other and
therefore not dependent on the way in which the pyrene-
modified UNA monomer was incorporated into the oligonu-
cleotide backbone. Generally, the DNA*/DNA duplexes were
more stable than the DNA*/RNA duplexes; this indicates that
the pyrene moiety fits better into the major groove of a DNA/
DNA duplex (see also UV/visible absorption studies). Even
though duplexes containing pyrene-modified UNA monomers
were found to be less stable than the unmodified references,
The thermal stabilities of duplexes containing monomers X
and Y incorporated as single-nucleotide bulges were also stud-
ied (Table 3). DNA*/DNA duplexes with single bulge-incorporat-
ed X or Y (ON1 and ON4) showed Tm values similar to those of
the unmodified fully matched dsDNAs, whereas the corre-
Table 1. Thermal denaturation temperatures (Tm)[a] of matched and mismatched duplexes based on complementary DNA and RNA strands with mismatch-
es in the central position 11 of the complementary strand.
Code
DNA/RNA[b]
Sequence
3’-ACG TGA CAT ABA GAC ATG GTA
B (DNA)
B (RNA)
C
A
G
T
C
A
G
U
DNA
ON1
ON2
ON3
ON4
ON5
ON6
5’-TGC ACT GTA TGT CTG TAC CAT
5’-TGC ACT GTA XGT CTG TAC CAT
5’-TGC ACT GTA XGX CTG TAC CAT
5’-TGC ACX GTA TGT CTG XAC CAT
5’-TGC ACT GTA YGT CTG TAC CAT
5’-TGC ACT GTA YGY CTG TAC CAT
5’-TGC ACY GTA TGT CTG YAC CAT
63.1
58.9
52.0
55.9
58.9
51.7
55.9
[À10.1]
[À8.0]
[À2.5]
[À12.3]
[À8.2]
[À2.4]
[À12.0]
[À8.6]
[À7.6]
[À2.3]
[À9.8]
[À8.2]
[À1.6]
[À9.8]
[À7.5]
[À6.4]
[À2.3]
[À8.9]
[À5.5]
[À2.0]
[À8.7]
63.3
57.7
48.5
51.7
58.3
49.8
52.2
[À10.4]
[À8.1]
[0.0]
[À12.3]
[À8.5]
[À0.2]
[À12.4]
[À8.2]
[À6.8]
[+0.7]
[À9.0]
[À4.9]
[À0.3]
[À9.7]
[À7.9]
[À5.0]
[À0.4]
[À9.4]
[À8.7]
[À1.2]
[À9.6]
[a] Tm values (8C) were measured as the maxima of the first derivatives of the thermal melting curves (A260 vs. temperature) recorded in medium-salt buffer
(100 mm NaCl, 0.1 mm EDTA, 10 mm NaH2PO4, 5 mm Na2HPO4, pH 7.0) with use of 1.0 mm concentrations of the two strands. DTm values for mismatches
are given in brackets. DTm values were calculated relative to fully matched duplexes of studied ONs. [b] In RNA targets, uracil bases are present instead of
thymine bases.
ꢂ 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemBioChem 2014, 15, 146 – 156 148

CHEMBIOCHEM
FULL PAPERS
www.chembiochem.org
Table 2. Thermal denaturation temperatures (Tm)[a] of matched and mismatched duplexes based on complementary DNA and RNA strands with mismatch-
es directly opposite the modification (position 10).
Code
DNA/RNA[b]
Sequence
3’-ACG TGA CAT BCA GAC ATG GTA
B (DNA)
B (RNA)
A
G
T
C
A
G
U
C
DNA
ON1
ON2
ON3
ON4
ON5
ON6
5’-TGC ACT GTA TGT CTG TAC CAT
5’-TGC ACT GTA XGT CTG TAC CAT
5’-TGC ACT GTA XGX CTG TAC CAT
5’-TGC ACX GTA TGT CTG XAC CAT
5’-TGC ACT GTA YGT CTG TAC CAT
5’-TGC ACT GTA YGY CTG TAC CAT
5’-TGC ACY GTA TGT CTG YAC CAT
63.1
58.9
52.0
55.9
58.9
51.7
55.9
[À5.7]
[+0.6]
[À0.4]
[À5.9]
[À0.8]
[À0.8]
[À6.6]
[À7.8]
[À9.4]
63.3
57.7
48.5
51.7
58.3
49.8
52.2
[À4.0]
[+0.8]
[À2.6]
[À5.5]
[À1.0]
[À2.3]
[À4.9]
[À7.6]
[À9.0]
[À1.7]
[À2.1]
[À9.1]
[À2.1]
[À2.6]
[À9.5]
[À2.6]
[À1.0]
[À11.4]
[À2.0]
[À0.7]
[À12.9]
[À0.8]
[À0.6]
[À8.3]
[À1.3]
[À0.6]
[À8.7]
[À2.2]
[À1.8]
[À12.6]
[À2.6]
[À1.9]
[À14.4]
[a], [b] See the footnotes of Table 1.
UNA monomer U (UNA1) units were both found to be approxi-
mately 48C less stable than the corresponding unmodified
fully matched duplexes.
Table 3. Thermal denaturation temperatures (Tm) of matched duplexes
based on complementary DNA and RNA strands and of duplexes incorpo-
rating bulges.
Introduction of two bulge-incorporations separated by one
base (UNA2, ON2 and ON5) had a destabilizing effect on both
DNA*/DNA and DNA*/RNA duplexes, with the duplex stabilities
following the order Y>X@U with respect to the introduced
modifications.
Tm [8C][a]
Code
Sequence
DNA
RNA
DNA/RNA[b]
DNA
3’-ACG TGA CAT –CA GAC ATG GTA
5’-TGC ACT GTA –GT CTG TAC CAT
5’-TGC ACT GTA TGT CTG TAC CAT
61.7
63.3
DNA bulge
57.4
[À4.3]
58.2
59.2
[À4.1]
59.3
Finally, thermal stabilities of nucleic acid duplexes containing
two bulge-incorporations separated by nine bases (UNA3, ON3
and ON6) were studied. The presence of two bulge-incorporat-
ed UNA monomers U (UNA3) led to significant destabilization
of the duplexes. In contrast, DNA*/DNA and DNA*/RNA duplex-
es containing two bulge-incorporated monomers X (ON3)
showed only minor thermal destabilization. The presence of
two bulge-incorporated monomers Y separated by nine bases
(ON5) led to stabilization of the DNA*/DNA duplex by 28C and
to destabilization of the DNA*/RNA duplex by 38C relative to
the fully matched unmodified duplexes. The data also showed
that generally the Tm values for the duplexes containing bulge-
incorporated monomers X and Y were higher for DNA*/DNA
duplexes than for DNA*/RNA duplexes, thus indicating that
bulge-incorporated UNA monomers with the pyrene-modified
nucleobase are better able to adapt to the conformation of
a DNA/DNA duplex than to that of a DNA/RNA duplex. Further-
more, bulge-containing duplexes containing pyrene-modified
UNA monomers were always significantly more stable than un-
modified duplexes containing DNA-T bulges. It is also impor-
tant to mention that duplexes with bulge-incorporated mono-
mers X and Y were 2–108C more stable than fully matched du-
plexes of ON1–6 with complementary strands (Table 1).
The substantial differences between the stabilities of
matched and bulge-containing duplexes might be exploitable
for the development of antisense oligonucleotides and hybridi-
zation probes and for strand invasion studies. The increased
stabilities of the bulge-containing duplexes are presumably
due to intercalation of the pyrenyl moiety into the duplex (see
also UV/visible absorption studies). A B-type duplex is less
compressed and has more efficient nucleobase overlap than
an A-type duplex;[15] this is why the pyrene moiety is able to
intercalate favourably into a DNA/DNA duplex, resulting in
higher thermal stabilities of DNA*/DNA duplexes containing
UNA1
ON1
5’-TGC ACT GTA UGT CTG TAC CAT
5’-TGC ACT GTA XGT CTG TAC CAT
5’-TGC ACT GTA YGT CTG TAC CAT
[À3.5]
61.9
[À4.0]
61.9
[+0.2]
61.8
[À1.4]
60.5
ON4
[+0.1]
[À2.8]
DNA/RNA[b]
DNA
3’-ACG TGA CAT –C– GAC ATG GTA
5’-TGC ACT GTA –G– CTG TAC CAT
5’-TGC ACT GTA TGT CTG TAC CAT
62.2
49.4
62.1
51.7
DNA bulge
[À12.8]
47.8
[À10.4]
50.6
UNA2
ON2
5’-TGC ACT GTA UGU CTG TAC CAT
5’-TGC ACT GTA XGX CTG TAC CAT
5’-TGC ACT GTA YGY CTG TAC CAT
[À14.4]
55.0
[À11.5]
52.8
[À7.2]
58.1
[À9.3]
53.7
ON5
[À4.1]
[À8.4]
DNA/RNA[b]
DNA
3’-ACG TG– CAT ACA GAC –TG GTA
5’-TGC AC– GTA TGT CTG –AC CAT
5’-TGC ACT GTA TGT CTG TAC CAT
63.6
50.2
63.4
51.8
DNA bulge
[À13.4]
50.7
[À11.6]
53.0
UNA3
ON3
5’-TGC ACU GTA TGT CTG UAC CAT
5’-TGC ACX GTA TGT CTG XAC CAT
5’-TGC ACY GTA TGT CTG YAC CAT
[À13.6]
62.6
[À10.4]
62.1
[À1.0]
65.4
[À1.3]
60.3
ON6
[+1.8]
[À3.1]
[a] Tm values (8C) were measured as the maxima of the first derivatives of
the thermal melting curves (A260 vs. temperature) recorded in medium-
salt buffer (100 mm NaCl, 0.1 mm EDTA, 10 mm NaH2PO4, 5 mm Na2HPO4,
pH 7.0) with 1.0 mm concentrations of the two strands. DTm values for du-
plexes containing bulges are given in brackets. Unmodified fully matched
DNA/DNA or DNA/RNA duplexes were used as references. U=uracil-1-yl
UNA monomer. [b] In RNA targets, uracil bases are present instead of thy-
mine bases.
sponding DNA*/RNA duplexes were 1–38C less stable than the
unmodified fully matched DNA/RNA duplexes. On the other
hand, the two duplexes containing single bulge-incorporated
ꢂ 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemBioChem 2014, 15, 146 – 156 149

CHEMBIOCHEM
FULL PAPERS
www.chembiochem.org
shown to induce an i-motif structure.[27] Nevertheless, destabili-
zation effects of monomers X or Y were still observed. General-
ly, the destabilization effect was higher for 3’-linked monomer
X than for 2’-linked monomer Y. The backbone of monomer Y
is two atoms longer than the that of monomer X, and the 2’-
linked monomer Y can therefore introduce a higher degree of
flexibility into the loop region; this possibly makes the i-motif
structure more stable than with the 3’-linked monomer X. This
is in agreement with the previous observations that an in-
creased level of flexibility in the loop regions (positions 4, 10–
12 and 16) is favourable for i-motif stability.[7] On the other
hand, the presence of the pyrene moiety in UNA monomers X
and Y led to lower stability of the i-motif structures relative to
i-motifs with unmodified UNA incorporations,[7] presumably
due to increased stacking interactions with neighbouring nu-
cleobases leading to structural perturbations of the i-motif
loop region, as also indicated by molecular modelling (see the
Supporting Information).
Table 4. Thermal denaturation temperatures (Tm) of DNA duplexes con-
taining bulges and effects of the central mismatch (position 11 in ON6).
Sequence (code)
T
m [8C][a]
3’-ACG TG– CAT ABA GAC –TG GTA
5’-TGC AC– GTA TGT CTG –AC CAT (DNA)
5’-TGC ACT GTA TGT CTG TAC CAT
(DNA bulge)
B=C [A; G; T]
63.6 [À10.7; À8.7; À8.0]
50.2 [À12.4; À11.0; À9.9]
5’-TGC ACY GTA TGT CTG YAC CAT (ON6)
65.4 [À9.9; 7.2; À7.2]
[a] Tm values (8C) were measured as the maxima of the first derivatives of
the thermal melting curves (A260 vs. temperature) recorded in medium-
salt buffer (100 mm NaCl, 0.1 mm EDTA, 10 mm NaH2PO4, 5 mm Na2HPO4,
pH 7.0) with 1.0 mm concentrations of the two complementary strands.
DTm values for mismatches are given in brackets in the following order:
G:A mismatch, G:G mismatch, G:T mismatch. DTm values were calculated
relative to matched duplexes of studied ONs.
monomers X and Y than in the case of the corresponding
DNA*/RNA duplexes.
Because increased thermal stability of the DNA*/DNA duplex
containing two bulge-incorporated monomers Y (ON6) was
observed, we decided to study the mismatch discrimination of
the central mismatch (position 11 in ON6) for this ON (Table 4).
The mismatch discrimination capabilities were found to be
slightly lower than those of the unmodified 19-mer dsDNA,
but sufficient for application as a probe for, for example, nucle-
ic acid detection. A DNA/DNA duplex containing two DNA-T
bulges showed high mismatch discrimination (Table 4), thus
supporting the trend of destabilization of the terminal parts of
the duplex resulting in improved mismatch sensing in the cen-
tral region.
Steady-state fluorescence emission studies
The fluorescence properties of oligonucleotides ON1 and ON4,
of the corresponding matched duplexes and of duplexes con-
taining single bulges were studied. Steady-state fluorescence
emission spectra were recorded at an excitation wavelength of
350 nm. Spectra of single-stranded ON1, of duplexes formed
between ON1 and complementary DNA and RNA strands and
of duplexes of ON1 with single-bulge incorporated mono-
mer X showed broad emission maxima around 452 nm (Fig-
ure 1A). The fluorescence intensities were similar for single-
stranded oligonucleotide ON1 and its duplex with complemen-
tary DNA, whereas an approximately 1.5-fold increase in fluo-
rescence intensity relative to single-stranded ON1 was ob-
served for the bulge-containing DNA*/DNA duplex of ON1.
The fluorescence intensities of the matched DNA*/RNA duplex
of ON1 and the DNA*/RNA duplex with a bulge-incorporated
monomer X were found to be lower than the fluorescence
intensity of single-stranded ON1. The fluorescence intensity of
the bulge-containing duplex was slightly higher than that of
the fully matched duplex.
Monomers X and Y were incorporated into position 10 (loop
region) in the C-rich 22 nt fragment of human telomeric DNA
(ON7, ON8, Table 5). The loop regions of ON7 and ON8 (XAA
Table 5. Thermal denaturation temperatures (Tm)[a] of DNA i-motifs.
Conditions
Sequence (code)
Noncrowding Crowding
5’-CCC TAA CCC TAA CCC TAA CCC T (wt)
5’-CCC TAA CCC XAA CCC TAA CCC T (ON7)
5’-CCC TAA CCC YAA CCC TAA CCC T (ON8)
45.6
39.7
43.3
41.1
37.9
38.3
The emission maxima of ON4, of its duplexes with comple-
mentary DNA and RNA strands and of its duplexes with bulge-
incorporated monomer Y were observed at approximately
452 nm (Figure 1B). Although the fluorescence intensities of
single-stranded ON1 and ON4 were similar, the intensities of
all duplexes of ON4 were significantly lower than that of the
single-strand ON4. The bulge-containing DNA*/DNA duplex of
ON4 showed higher fluorescence intensity than the fully
matched duplex, following the same trend as observed with
ON1. On the other hand, both the bulge-containing DNA*/
RNA duplex of ON4 and the matched duplex of ON4 with RNA
were found to display similar fluorescence intensities. The
observed emission maxima of oligonucleotides ON1 and ON4
and of their duplexes originate from an exciplex involving the
5-(pyren-1-yl)uracil unit in monomers X and Y, which is partially
incorporated into the nucleobase stack. Similar results were
previously reported with DNA duplexes containing a 5-(pyren-
[a] Tm values (8C) were measured as the minima of the first derivatives of
the thermal melting curves (A295 vs. temperature) recorded with 3.0 mm
concentrations of the strands. Noncrowding conditions: 100 mm KCl,
10 mm sodium cacodylate, pH 5.2. Crowding conditions: 100 mm KCl,
10 mm sodium cacodylate, 20% (v/v) PEG 200, pH 5.2.
and YAA, respectively) are therefore closely structurally related
to the native sequence (TAA). The Tm values of the i-motif
structures formed by these ONs were studied (Table 5): both
monomers X and Y were found to destabilize the i-motifs at
pH 5.2. The Tm values determined under molecular crowding
conditions (20% PEG 200), mimicking an intracellular environ-
ment, were lower than those under noncrowding conditions,
despite the fact that crowding conditions had previously been
ꢂ 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemBioChem 2014, 15, 146 – 156 150

CHEMBIOCHEM
FULL PAPERS
www.chembiochem.org
Figure 1. A) Steady-state fluorescence emission spectra of ON1 in matched duplexes, duplexes containing single bulges and as a single strand. B) Steady-
state fluorescence emission spectra of ON4 in matched duplexes, duplexes containing single bulges and as a single strand. C) Absorption spectrum of ON1
in matched duplexes, duplexes containing single bulges and as a single strand. D) Absorption spectrum of ON4 in matched duplexes, duplexes containing
single bulges and as a single strand. Measurements were obtained with 1.0 mm concentrations of the two strands in medium-salt buffer (100 mm NaCl,
0.1 mm EDTA, 10 mm NaH2PO4, 5 mm Na2HPO4, pH 7.0) at 208C. lex =350 nm.
1-yl)-2’-deoxyuridine unit.[21,23] The variations observed in the
fluorescence intensities are probably due to changes in the p-
stacking interactions and the electron-transfer reactions be-
tween monomers X and Y and neighbouring nucleobases. The
fluorescence intensities of ON1 and ON4 were substantially
lower than those of ONs containing the 2’-pyrene-modified
UNA monomers W and Z.[6] On the other hand, due to the
direct conjugation of pyrene with a nucleobase, the emission
maxima of ON1 and ON4 each showed a bathochromic shift
of approximately 180 nm in relation to ONs containing 2’-
pyrene-modified UNAs.[6] Larger Stokes shifts of 5-pyrene-modi-
fied UNAs might be beneficial for, for example, fluorescence
imaging measurements.
480 nm and similar fluorescence intensities (Figure 2A and B).
The emission maximum of single-stranded ON7 at pH 7.0
(457 nm) was blue-shifted in relation to that of the matched
duplex, whereas its fluorescence intensity was halved. Efficient
quenching of fluorescence was observed for ON7 at pH 5.2
(Figure 2A): the fluorescence of ON7 at pH 5.2 at 457 nm was
quenched by 89% relative to the fluorescence of ON7 at
pH 7.0.
The spectrum of ON8 at pH 7.0 showed a broad maximum
at about 460 nm, whereas at pH 5.2 the fluorescence of ON8
at 460 nm was quenched by 90% (Figure 2B). It follows that
the formation of an i-motif structure under acidic conditions
quenches the fluorescence of ONs with 5-(pyren-1-yl)uracilyl
UNA monomers in the loop region. One of the possible ex-
planations for this fact is p-stacking interactions or electron-
transfer reactions with the neighbouring nucleobases (see mo-
lecular modelling in the Supporting Information). Although
changes in fluorescence intensities of ONs containing fluores-
Next, fluorescence emission spectra of the i-motif-forming
oligonucleotides ON7 and ON8 and of their matched DNA*/
DNA duplexes were recorded at 350 nm excitation wavelength.
The matched duplexes of ON7 and ON8 with complementary
DNA showed broad emission maxima between 450 and
ꢂ 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemBioChem 2014, 15, 146 – 156 151

CHEMBIOCHEM
FULL PAPERS
www.chembiochem.org
Figure 2. A) Steady-state fluorescence emission spectra of ON7 in matched duplex (pH 7.0) and as a single strand (pH 7.0 and 5.2). B) Steady-state fluores-
cence emission spectra of ON8 in matched duplex (pH 7.0) and as a single strand (pH 7.0 and 5.2). C) Absorption spectrum of ON7 in matched duplex
(pH 7.0) and as a single strand (pH 7.0 and 5.2). D) Absorption spectra of ON8 in matched duplex (pH 7.0) and as a single strand (pH 7.0 and 5.2). Measure-
ments were obtained with 1.0 mm concentrations of the strands in cacodylate buffer (100 mm KCl, 10 mm sodium cacodylate) at 208C. Excitation wavelength:
350 nm.
cently labelled nucleobases have been reported upon forma-
tion of an i-motif structure,[24,28] such efficient quenching of the
fluorescence has not previously been observed. Such a dramat-
ic change in fluorescence properties might be caused by
strong electronic coupling between the nucleobase and the
pyrene moiety in monomers X and Y, because the two chro-
mophores are linked covalently by a single CÀC bond, whereas
in the previously reported nucleobase-modified ONs the fluo-
rescent label was attached through an ethylene linker. The effi-
cient quenching of fluorescence upon formation of an i-motif
structure might be useful for probing such i-motif structures.
strands and of duplexes containing bulge-incorporated mono-
mers X and Y were recorded (310–410 nm). ON1 showed an
absorption maximum at 350 nm, as typically observed for a
pyrenyl group (Figure 1C). Shoulders at 335 and 356 nm were
also observed. The duplexes of ON1 with complementary DNA
and RNA both showed slightly broader absorptions in the
340–360 nm region, and the A350/A356 ratios were slightly de-
creased, thus indicating small bathochromic shifts in the ab-
sorption spectra, presumably due to partial intercalation of the
uracil part of the 5-(pyren-1-yl)uracil moiety into the base
stack. The DNA*/DNA duplex of ON1 containing bulge-incor-
porated monomer X showed two local absorption maxima at
345 and 360 nm. Notably, the 10 nm shift of the absorption
maximum of the pyrene unit to 360 nm indicates that the
pyrene moiety in the bulge intercalates into the DNA*/DNA
duplex. Local absorption maxima of the DNA*/RNA duplex of
ON1 containing bulge-incorporated monomer X were located
at 347 and 356 nm. The bathochromic shift of the pyrene ab-
UV/visible absorption studies
In the next part of our study we focused on UV/visible absorp-
tion studies of the ONs containing pyrene-modified UNA mon-
omers X and Y. The UV/visible spectra of oligonucleotides ON1
and ON4, of their duplexes with complementary DNA and RNA
ꢂ 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemBioChem 2014, 15, 146 – 156 152

CHEMBIOCHEM
FULL PAPERS
www.chembiochem.org
sorption maximum in the bulge-containing DNA*/RNA duplex
was less profound than in the DNA*/DNA duplex, presumably
due to the fact that the intercalation into DNA/RNA duplex is
more demanding and less favourable.
DNA/RNA duplexes. The discovery that monomer Y can have
either stabilizing or destabilizing effects in nucleic acid duplex-
es, depending on the way in which it is incorporated, might
be useful for the design of antisense ONs and hybridization
probes. Steady-state fluorescence emission studies of ONs with
single incorporation of pyrene-modified monomers X and Y in
most cases revealed decreases in fluorescence intensity upon
hybridization to DNA or RNA. Efficient quenching of the fluo-
rescence of i-motif-forming ONs containing pyrene-modified
UNA monomers was observed at pH 5.2. To the best of our
knowledge this is so far the most effective fluorescence
quenching upon formation of an i-motif structure.
The same trend was observed when the UV/visible absorp-
tion of ON4 and of its duplexes were studied. Single-stranded
ON4 and duplexes of ON4 with complementary DNA and RNA
showed absorption maxima at approximately 353 nm (Fig-
ure 1D). On the other hand, absorption maxima and duplexes
of ON4 containing bulges were found to be around 345 and
360 nm, thus implying significant bathochromic shifts.
In summary, these results indicate that bulge-incorporated
UNA monomers containing pyrene-modified nucleobases are
able to intercalate into both DNA/DNA and DNA/RNA duplex-
es, with the pyrene moieties being located in the major
grooves in the matched duplexes of two ONs of identical nu-
cleotide length.
Experimental Section
NMR spectra were recorded with a 400 MHz (1H at 400 MHz, 13C at
110.6 MHz, 31P at 162 MHz) spectrometer in CDCl3 solutions. Chemi-
cal shifts are reported in ppm relative to TMS (1H, internal standard,
dH TMS=0.00 ppm), solvent peaks (13C, internal standard, dC
CDCl3 =77.00 ppm), or 85% H3PO4 (31P, external standard, dp 85%
H3PO4 =0.00 ppm). Assignments of the NMR signals are based on
2D correlation experiments. High-resolution MS spectra were col-
lected by use of electrospray ionization. ONs were synthesized
with an ꢃKTA oligopilot Plus system (GE Healthcare Life Sciences),
and the compositions of ONs were confirmed by MALDI-MS (Micro-
flex MALDI-TOF MS, Bruker Daltonics) and by their purities (>90%)
by ion-exchange HPLC.
The UV/visible absorption properties of the i-motif-forming
oligonucleotides ON7 and ON8 were also studied. At pH 7.0
ON7 and ON8 each showed the typical absorption maximum
for single-stranded ONs containing pyrene-modified UNA mon-
omers at approximately 350 nm (Figure 2C, 2D). At pH 5.2,
however, small bathochromic shifts of the absorption maxima
of ON7 and ON8 were observed. These shifts indicate stacking
interactions between the pyrene-modified nucleobases and
the neighbouring nucleobases when i-motif structures are
present at pH 5.2. More intensive stacking was indicated for
the 3’-linked monomer X, presumably due to the expected
lower flexibility of the loop region of the i-motif for ON7 (con-
taining monomer X) relative to ON8.
5’-O-(4,4’-Dimethoxytrityl)-5-iodo-2’,3’-secouridine (2): 5-Iodo-
uridine[25] (1, 3.15 g, 4.69 mmol) was dissolved in a mixture of 1,4-
dioxane (60 mL) and water (12 mL). A solution of NaIO4 (1.10 g,
5.16 mmol) in water (10 mL) was added, and the reaction mixture
was stirred for 1 h at RT. 1,4-Dioxane (50 mL) was then added, and
stirring was continued for 15 min. The precipitate was filtered off
and washed with 1,4-dioxane (25 mL). NaBH4 (203 mg, 5.34 mmol)
was added to the filtrate, and the mixture was stirred for 1 h at RT.
A mixture of acetic acid and pyridine (6 mL, 1:1, v/v) was then
added, and the resulting mixture was concentrated under reduced
pressure to a volume of approximately 25 mL. CH2Cl2 (50 mL) was
added to this solution, and washing was performed with saturated
aqueous NaHCO3 (50 mL). The separated organic phase was dried
over MgSO4 and, after filtration, concentrated to dryness under re-
duced pressure. The residue was purified by silica gel column chro-
matography (0–2.5% MeOH in CH2Cl2, v/v) to furnish nucleoside 2
Conclusions
We have developed a synthesis of two new phosphoramidites
suitable for incorporation of 5-(pyren-1-yl)uracilyl UNA mono-
mers X and Y into ONs. Monomers X and Y were found to
destabilize i-motif structures at pH 5.2, both under molecular
crowding and noncrowding conditions. The presence of one
or two of these pyrene-modified UNA monomers in a DNA
strand led to decreases in the thermal stabilities of DNA*/DNA
and DNA*/RNA duplexes, but the duplex thermal stabilities
were higher than those of previously reported[6] duplexes con-
taining unmodified UNA monomers (U), presumably due to
major groove interactions of the pyrene moiety. The mismatch
discrimination of ON1–6 varied from low to high depending
on the relative positioning of monomers within the duplexes.
The presence of two bulge-incorporated pyrene-modified UNA
monomers Y (nine bases apart) was able to stabilize a DNA*/
DNA duplex relative to the unmodified matched duplex. In
general, however, the thermal stabilities of DNA*/DNA and
DNA*/RNA duplexes containing bulge-incorporated pyrene-
modified monomers were similar to or slightly lower than
those of unmodified matched duplexes, but thus significantly
better than those of matched duplexes containing X and Y
monomers. From UV/visible absorption studies we conclude
that bulge-incorporated UNA monomers containing pyrene-
modified nucleobases intercalate into both DNA/DNA and
1
(2.28 g, 72%) as a white foam. H NMR (400 MHz, CDCl3): d=3.08
(dd, Jgem =10.4 Hz, J5’a,4’ =6.1 Hz, 1H; H-5’a), 3.18 (dd, Jgem =10.4 Hz,
J
5’b,4’ =3.7 Hz, 1H; H-5’b), 3.64–3.82 (m, 11H; H-2’,3’,4’, OCH3), 5.95
(t, J1’,2’ =5.3 Hz, 1H; H-1’), 6.74–6.82 (m, 4H; H-3-PhOMe), 7.12–7.27
(m, 7H; H-o,p-Ph, H-2-PhOMe), 7.29–7.35 (m, 2H; H-m-Ph),
7.79 ppm (s, 1H; H-6); 13C NMR (100.6 MHz, CDCl3): d=55.24
(OCH3), 62.90, 63.49, 63.65 (CH2-2’, CH2-3’, CH2-5’), 69.18 (C-5), 81.83
(CH-4’), 85.93 (CH-1’), 86.58 (CPh(PhOMe)2), 113.25 (CH-3-PhOMe),
126.92 (CH-p-Ph), 127.92 (CH-o,m-Ph), 129.84 and 129.89 (CH-2-
PhOMe), 135.41 and 135.54 (C-1-PhOMe), 144.31 (C-i-Ph), 144.90
(CH-6), 150.92 (C-2), 158.47 (C-4-PhOMe), 160.51 ppm (C-4); HRMS
(ESI): m/z calcd for C30H31IN2NaO8: 697.1017 [M+Na]+; found:
697.0994.
2’-O-Benzoyl-5’-O-(4,4’-dimethoxytrityl)-5-iodo-2’,3’-secouridine
(3): A solution of nucleoside 2 (645 mg, 0.95 mmol) and DBU
(286 mL, 1.91 mmol) in anhydrous CH2Cl2 (25 mL) was cooled to
À788C, and a solution of benzoyl chloride in CH2Cl2 (0.5m,
2.00 mL, 1.00 mmol) was added dropwise over a period of 20 min.
ꢂ 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemBioChem 2014, 15, 146 – 156 153

CHEMBIOCHEM
FULL PAPERS
www.chembiochem.org
The reaction mixture was stirred at À788C for 30 min. MeOH
(0.9 mL) was then added, and the mixture was allowed to warm to
RT. The mixture was extracted with saturated aqueous NaHCO3
(50 mL). The separated organic phase was dried over MgSO4 and,
after filtration, was concentrated to dryness under reduced pres-
sure. The crude product was purified by high-performance FLASH
chromatography (HPFC) on SiO2 (0–53% EtOAc in petroleum ether)
to furnish benzoate 3 (522 mg, 70%) as a white foam. 1H NMR
(400 MHz, CDCl3): d=3.12 (dd, Jgem =10.6 Hz, J5’a,4’ =6.0 Hz, 1H; H-
5’a), 3.23 (dd, Jgem =10.6 Hz, J5’b,4’ =4.0 Hz, 1H; H-5’b), 3.69–3.82 (m,
9H; H-3’,4’, OCH3), 4.44 (dd, Jgem =11.9 Hz, J2’a,1’ =4.5 Hz, 1H; H-2’a),
added, and stirring was continued for another 2 h. The reaction
was quenched by addition of MeOH (0.5 mL), and the resulting
mixture was stirred for another 5 min. The solution was diluted
with CH2Cl2 (10 mL) and washed with saturated aqueous NaHCO3
(15 mL). The organic phase was dried over MgSO4 and, after filtra-
tion, was concentrated to dryness under reduced pressure. The res-
idue was dissolved in EtOAc (2 mL) and added dropwise to ice-
cold petroleum ether (100 mL). The precipitate was filtered off and
dried under vacuum. Phosphoramidite 5 (163 mg, 93%) was ob-
tained as an off-white solid. 31P NMR (162 MHz, CDCl3): d=148.65
and 149.06 ppm; HRMS (ESI): m/z calcd for C62H61N4NaO10P:
1075.4018 [M+Na]+; found: 1075.3987.
4.63 (dd, Jgem =11.9 Hz, J2’b,1’ =4.5 Hz, 1H; H-2’b), 6.20 (t, J1’,2’
=
4.5 Hz, 1H; H-1’), 6.77–6.81 (m, 4H; H-3-PhOMe), 7.15–7.33 (m, 7H;
H-o,p-Ph, H-2-PhOMe), 7.30–7.34 (m, 2H; H-m-Ph), 7.43–7.48 (m,
2H; H-m-Bz), 7.57–7.61 (m, 1H; H-p-Bz), 7.84 (s, 1H; H-6), 7.96–8.00
(m, 2H; H-o-Bz), 8.95 ppm (brs, 1H; NH); 13C NMR (100.6 MHz,
CDCl3): d=55.23 (OCH3), 62.38 (CH2-5’), 63.39 (CH2-3’), 64.21 (CH2-
2’), 68.72 (C-5), 82.42 (CH-4’), 83.30 (CH-1’), 86.73 (CPh(PhOMe)2),
113.27 (CH-3-PhOMe), 126.95 (CH-p-Ph), 127.88 (CH-Ar), 127.97 (CH-
Ar), 128.71 (CH-Ar), 128.82 (CH-Ar), 129.76 (CH-Ar), 129.82 (CH-Ar),
129.87 (CH-Ar), 133.69 (CH-p-Bz), 135.24 and 135.39 (C-1-PhOMe),
144.21 (C-i-Ph), 144.39 (CH-6), 149.91 (C-2), 158.53 (C-4-PhOMe),
159.67 (C-4), 171.17 ppm (C=O benzoyl). HRMS (ESI): m/z calcd for
C37H35IN2NaO9: 801.1279 [M+Na]+; found: 801.1288.
3’-O-(tert-Butyldimethylsilyl)-5’-O-(4,4’-dimethoxytrityl)-5-iodo-
2’,3’-secouridine (6): Nucleoside 3 (511 mg, 0.66 mmol) was co-
evaporated with anhydrous pyridine (2ꢄ5 mL). The residue was
dissolved in anhydrous pyridine (10 mL), and TBSCl (593 mg,
3.94 mmol) was added. The reaction mixture was stirred at RT over-
night. Water (5 mL) was then added, and stirring was continued for
20 min at RT. The resulting mixture was diluted with MeOH
(50 mL), after which sodium hydroxide (735 mg, 18.37 mmol) was
added. The reaction mixture was stirred at RT for 2 h. The mixture
was then diluted with CH2Cl2 (30 mL) and washed with saturated
aqueous NH4Cl (30 mL). The organic phase was dried over MgSO4
and, after filtration, was concentrated to dryness under reduced
pressure. Purification by silica gel column chromatography (20–
30% EtOAc in petroleum ether) afforded compound 6 (394 mg,
2’-O-Benzoyl-5’-O-(4,4’-dimethoxytrityl)-5-(pyren-1-yl)-2’,3’-se-
couridine (4): A nitrogen-purged mixture of benzoate 3 (150 mg,
0.19 mmol), pyrene-1-boronic acid (142 mg, 0.58 mmol), sodium
carbonate (61 mg, 0.58 mmol), PPh3 (10 mg, 39 mmol) and Pd(OAc)2
(4.3 mg, 19 mmol) in toluene (4.5 mL) and water (675 mL) was
stirred at 1008C for 1 h. After the system had cooled, volatiles
were removed under reduced pressure. The residue was dissolved
in CH2Cl2 (20 mL) and washed with water (30 mL). The organic
phase was dried over MgSO4 and, after filtration, was concentrated
to dryness under reduced pressure. The residue was dissolved in
CH2Cl2 and coevaporated with silica gel. The product was purified
by silica gel column chromatography (0–1% MeOH in CH2Cl2) to
furnish compound 4 (58 mg, 35%) as a yellowish amorphous solid.
1H NMR (400 MHz, CDCl3): d=3.14 (dd, Jgem =10.2 Hz, J5’a,4’ =6.0 Hz,
1H; H-5’a), 3.26 (dd, Jgem =10.2 Hz, J5’b,4’ =5.0 Hz, 1H; H-5’b), 3.51,
3.54 (2s, 3H; OCH3), 3.64–3.73 (m, 1H; H-3’a), 3.78–3.86 (m, 1H; H-
3’b), 3.91–3.98 (m, 1H; H-4’), 4.57 (dd, Jgem =11.8 Hz, J2’a,1’ =4.6 Hz,
1H; H-2’a), 4.75 (dd, Jgem =11.8 Hz, J2’b,1’ =4.9 Hz, 1H; H-2’b), 6.38 (t,
1
76%) as a white foam. H NMR (400 MHz, CDCl3): d=0.06 (s, 6H;
2SiCH3), 0.88 (s, 9H; C(CH3)3), 3.09 (dd, Jgem =10.6 Hz, J5’a,4’ =6.1 Hz,
1H; H-5’a), 3.18 (dd, Jgem =10.5 Hz, J5’b,4’ =3.9 Hz, 1H; H-5’b), 3.62–
3.78 (m, 5H; 2H-3’, 2H-2’, H-4’), 3.79 (s, 6H; 2OCH3), 5.90 (dd,
J
1’,2’a =5.8 Hz, J1’,2’b =4.0 Hz, 1H; H-1’), 6.79–6.83 (m, 4H; H-3-
PhOMe), 7.17–7.37 (m, 9H; H-Ph, H-2-PhOMe), 7.83 (s, 1H; H-6),
8.77 ppm (brs, 1H; NH); 13C NMR (100.6 MHz, CDCl3): d=À5.50
(SiCH3), 18.28 (C(CH3)3), 25.82 (C(CH3)3), 55.25 (OCH3), 63.46, 63.52,
63.92 (CH2-2’, CH2-3’, CH2-5’), 68.23 (C-5), 81.85 (CH-4’), 85.39 (CH-
1’), 86.64 (CPh(PhOMe)2), 113.24 (CH-3-PhOMe), 126.91 (CH-4-Ph),
127.92 (CH-2,3-Ph), 129.84 and 129.90 (CH-2-PhOMe), 135.50 and
135.61 (C-1-PhOMe), 144.39 (C-1-Ph), 144.93 (CH-6), 150.02 (C-2),
158.53 (C-4-PhOMe), 159.81 ppm (C-4); HRMS (ESI): m/z calcd for
C36H45IN2NaO8Si: 811.1882 [M+Na]+; found: 811.1914.
3’-O-(tert-Butyldimethylsilyl)-5’-O-(4,4’-dimethoxytrityl)-5-(pyren-
1-yl)-2’,3’-secouridine (7): Nucleoside 7 was prepared as described
for compound 4; nucleoside 6 (350 mg, 0.44 mmol), pyrene-1-bor-
onic acid (328 mg, 1.33 mmol), Pd(OAc)2 (10 mg, 44 mmol), PPh3
(23 mg, 89 mmol) and Na2CO3 (141 mg, 1.33 mmol) in toluene
(13 mL) and water (1.95 mL) were used. Purification by silica gel
column chromatography (0–0.75% MeOH in CH2Cl2) afforded nu-
J
1’,2’ =4.7 Hz, 1H; H-1’), 6.51–6.64 (m, 4H; H-3-PhOMe), 7.09–7.37
(m, 14H; H-Ph, H-2-PhOMe, H-Bz), 7.64 (s, 1H; H-6), 7.91–8.25 (m,
9H; H-pyrene), 8.78 ppm (brs, 1H; NH); 13C NMR (100.6 MHz,
CDCl3): d=54.99 (OCH3), 62.89 (CH2-3’), 63.25 (CH2-5’), 64.14 (CH2-
2’), 82.17 (CH-4’), 83.25 (CH-1’), 86.65 (CPh(PhOMe)2), 113.07 and
113.13 (CH-3-PhOMe), 124.17 (C-Ar), 124.57 (CH-Ar), 124.81 (C-Ar),
125.30 (CH-Ar), 125.41 (CH-Ar), 126.07 (CH-Ar), 126.89 (CH-Ar),
127.22 (CH-Ar), 127.67 (CH-Ar), 127.89 (CH-Ar), 128.19 (C-Ar), 128.59
(CH-Ar), 128.86 (C-Ar), 129.75 (CH-Ar), 129.79 (CH-Ar), 129.84 (CH-
Ar), 130.73 (C-Ar), 131.17 (C-Ar), 133.61 (C-Ar), 135.10 (C-Ar), 135.49
(C-Ar), 137.37 (CH-6), 144.56 (C-i-Ph), 150.20 (C-2), 158.46 (C-4-
PhOMe), 161.94 (C-4), 166.00 ppm (C=O benzoyl); HRMS (ESI): m/z
calcd for C53H44N2NaO9: 875.2939 [M+Na]+; found: 875.2966.
1
cleoside 7 (220 mg, 57%) as a yellowish amorphous solid. H NMR
(400 MHz, CDCl3): d=0.05 (s, 6H; 2SiCH3), 0.87 (s, 9H; C(CH3)3),
3.00–3.15 (m, 2H; H-5’a, 2’-OH), 3.20 (dd, Jgem =9.7 Hz, J5’b,4’
4.1 Hz, 1H; H-5’b), 3.50, 3.53 (2s, OCH3), 3.65 (dd, Jgem =10.8 Hz,
3’a,4’ =7.4 Hz, 1H; H-3’a), 3.68–4.00 (m, 4H; 2H-2’, H-3’b, H-4’), 6.09
=
J
(t, J1’,2’ =4.4 Hz, 1H; H-1’), 6.45–6.64 (m, 4H; H-3-PhOMe), 7.09–7.22
(m, 7H; DMT), 7.30–7.36 (m, 2H; DMT), 7.45 (td, J1 =7.4 Hz, J2 =
2.9 Hz, 1H; H-pyrene), 7.50–7.56 (m, 1H; H-pyrene), 7.60–7.71 (m,
3H; H-6, H-pyrene), 7.91–8.11 (m, 4H; H-pyrene), 8.16 (d, J=7.4 Hz,
1H; H-pyrene), 8.86 ppm (brs, 1H; NH); 13C NMR (100.6 MHz,
CDCl3): d=À5.49 (SiCH3), 18.29 (C(CH3)3), 25.82 (C(CH3)3), 54.97
(OCH3), 63.24 (CH2-5’), 63.57 (CH2-3’), 64.13 (CH2-2’), 82.07 (CH-4’),
85.58 (CH-1’), 86.50 (CPh(PhOMe)2), 113.01 and 113.06 (CH-3-
PhOMe), 124.27 (CH-Ar), 124.51 (CH-Ar), 124.57 (CH-Ar), 124.84 (CH-
Ar), 125.15 (CH-Ar), 125.23 (CH-Ar), 125.96 (CH-Ar), 126.82 (CH-Ar),
2’-O-Benzoyl-3’-O-[2-cyanoethoxy(diisopropylamino)phosphino]-
5’-O-(4,4’-dimethoxytrityl)-5-(pyren-1-yl)-2’,3’-secouridine (5): Nu-
cleoside 4 (142 mg, 0.17 mmol) was stirred with anhydrous DIPEA
(0.67 mmol) in anhydrous CH2Cl2 (10 mL), and 2-cyanoethyl-N,N-di-
isopropylchlorophosphoramidite [P(Cl)(NiPr2)(OCH2CH2CN), 56 mL,
0.25 mmol] was added. The mixture was stirred under N2 at RT for
2 h. Further P(Cl)(NiPr2)(OCH2CH2CN) (56 mL, 0.25 mmol) was then
ꢂ 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemBioChem 2014, 15, 146 – 156 154

CHEMBIOCHEM
FULL PAPERS
www.chembiochem.org
127.21 (CH-Ar), 127.67 (CH-Ar), 127.84 (CH-Ar), 128.34 (CH-Ar),
128.42 (CH-Ar), 128.54 (CH-Ar), 129.56 (CH-Ar), 129.79 (CH-Ar),
129.88 (CH-Ar), 130.72 (CH-Ar), 131.15 (CH-Ar), 131.90 (CH-Ar),
131.92 (CH-Ar), 132.04 (CH-Ar), 132.13 (CH-Ar), 135.26 (C-Ar), 135.66
(C-Ar), 140.30 (C-6), 144.73 (C-i-Ph), 150.32 (C-2), 158.42 ppm (C-4-
PhOMe); HRMS (ESI): m/z calcd for C52H54N2NaO8Si: 885.3542
[M+Na]+; found: 885.3518.
first derivatives of the thermal denaturation curves (A260 vs. T).
Reported Tm values are averages of two measurements within
Æ1.08C.
Thermal denaturation studies of i-motifs: Concentrations of ONs
were calculated by use of extinction coefficients,[29] including that
of the 5-(pyren-2-yl)uracil moiety.[23] ONs (3.0 mm) were mixed
either in sodium cacodylate buffer [KCl (100 mm), sodium cacody-
late (10 mm), pH 5.2] or in sodium cacodylate buffer with PEG 200
[KCl (100 mm), sodium cacodylate (10 mm), PEG 200 (20%, v/v),
pH 5.2], and the resulting mixtures were heated to 908C (10 min)
and then allowed to cool slowly to the starting temperature of the
experiment (158C). Thermal denaturation temperatures (Tm) were
determined with a Beckman DU 800 spectrophotometer as the
minima of the first derivatives of the thermal denaturation curves
(A295 vs. temperature). Reported Tm values are averages of two
measurements within Æ1.08C.
3’-O-(tert-Butyldimethylsilyl)-3’-O-[2-cyanoethoxy(diisopropyla-
mino)phosphino]-5’-O-(4,4’-dimethoxytrityl)-5-(pyren-1-yl)-2’,3’-
secouridine (8): Phosphoramidite 8 was prepared as described for
compound 5; nucleoside 7 (200 mg, 0.23 mmol), DIPEA (161 mL,
0.93 mmol) and P(Cl)(NiPr2)(OCH2CH2CN) (155 mL, 0.70 mmol) in
anhydrous CH2Cl2 (16 mL) were used. Phosphoramidite 8 (159 mg,
65%) was obtained as an off-white solid. 31P NMR (162 MHz,
CDCl3): d=149.47 ppm; HRMS (ESI): m/z calcd for C61H71N4NaO9PSi:
1085.4620 [M+Na]+; found: 1085.4614.
Steady-state fluorescence emission spectra: ONs were thorough-
ly mixed either in medium-salt buffer [NaCl (100 mm), EDTA
(0.1 mm), NaH2PO4 (10 mm), Na2HPO4 (5 mm), pH 7.0] or in sodium
cacodylate buffer [KCl (100 mm), sodium cacodylate (10 mm),
pH 5.2 or pH 7.0] in concentrations of 1 mm of each strand. The
samples were heated at 908C for 10 min and allowed to cool
slowly to 208C before measurements in quartz optical cells with
a path length of 1.0 cm. Background spectra of buffer solutions
were recorded with an excitation wavelength of 350 nm and were
subtracted from the relevant spectra. Steady-state spectra (360–
620 nm) were recorded with a PerkinElmer LS 55 luminescence
spectrometer at 208C as averages of three scans with use of an
excitation wavelength of 350 nm, an excitation slit of 4.0 nm, an
Oligonucleotide synthesis and purification: Synthesis of ONs was
performed on 1 mmol scale with an automated DNA synthesizer.
Standard cycle procedures were applied for unmodified phosphor-
amidites as well as for amidites 5 and 8 with use of a solution of
1H-tetrazole (0.45m) as an activator. Stepwise coupling yields, as
determined by a spectrophotometric DMT+ assay, were above
99% for phosphoramidites 5 and 8 (20 min coupling time) and un-
modified DNA phosphoramidites (2 min coupling time). Removal
of the nucleobase protecting groups and cleavage from solid sup-
port was effected under standard conditions (32% aqueous ammo-
nia, 12 h, 558C). Removal of the TBS group to obtain ON4–6 and
ON8 was accomplished by treatment with Et3N·3HF in N-methyl-2-
pyrrolidone in the presence of Et3N (4 h, 678C) and was followed
by detritylation (80% AcOH, 30 min, RT). ON1, ON4, ON7 and ON8
were precipitated from acetone after detritylation. ON2 and ON3
were purified by DMT-ON RP-HPLC with a C18-column, followed by
detritylation and acetone precipitation. ON5 and ON6 were puri-
fied by DMT-OFF ion-exchange HPLC, followed by acetone precipi-
tation. Analysis by ion-exchange HPLC verified the purity of all oli-
gonucleotides to be >90%, and their compositions were verified
by MALDI-TOF mass spectrometry (Table 6).
emission slit of 2.5 nm and a scan speed of 120 nmminÀ1
.
Acknowledgements
Funding from The University of Southern Denmark and The Euro-
pean Research Council under The European Union’s Seventh
Framework Programme (FP7/2007–2013)/ERC Grant agreement
No. 268776 is gratefully acknowledged.
Keywords: fluorescence · i-motifs · nucleic acid hybridization ·
oligonucleotides · unlocked nucleic acids
Table 6. MALDI-MS analysis of ON1–8.
Code
Sequence
Mcalcd
Mfound
ON1
ON2
ON3
ON4
ON5
ON6
ON7
ON8
5’-TGC ACT GTA XGT CTG TAC CAT
5’-TGC ACT GTA XGX CTG TAC CAT
5’-TGC ACX GTA TGT CTG XAC CAT
5’-TGC ACT GTA YGT CTG TAC CAT
5’-TGC ACT GTA YGY CTG TAC CAT
5’-TGC ACY GTA TGT CTG YAC CAT
5’-CCC TAA CCC XAA CCC TAA CCC T
5’-CCC TAA CCC YAA CCC TAA CCC T
6591.4
6795.6
6795.6
6591.4
6795.6
6795.6
6708.5
6708.5
6591.3
6797.0
6797.0
6592.2
6795.5
6799.8
6709.9
6712.2
[1] For reviews see: a) A. Pasternak, J. Wengel, Org. Biomol. Chem. 2011, 9,
3591–3597; b) H. Doessing, B. Vester, Molecules 2011, 16, 4511–4526;
c) M. A. Campbell, J. Wengel, Chem. Soc. Rev. 2011, 40, 5680–5689.
[2] P. Nielsen, L. H. Dreiøe, J. Wengel, Bioorg. Med. Chem. 1995, 3, 19–28.
[3] N. Langkjær, A. Pasternak, J. Wengel, Bioorg. Med. Chem. 2009, 17,
5420–5425.
[4] T. B. Jensen, N. Langkjær, J. Wengel, Nucleic Acids Symp. Ser. 2008, 52,
133–134.
[5] A. Pasternak, J. Wengel, Nucleic Acids Res. 2010, 38, 6697–6706.
[6] K. K. Karlsen, A. Pasternak, T. B. Jensen, J. Wengel, ChemBioChem 2012,
13, 590–601.
Thermal denaturation studies of nucleic acid duplexes: Concen-
trations of ONs were calculated by use of extinction coefficients,[29]
including that of the 5-(pyren-2-yl)uracil moiety.[23] ONs (1.0 mm of
each strand) were mixed in medium-salt buffer [NaCl (100 mm),
EDTA (0.1 mm), NaH2PO4 (10 mm), Na2HPO4 (5 mm), pH 7.0], and
the resulting mixtures were each heated to 908C (10 min) and al-
lowed to cool slowly to the starting temperature of the experiment
(208C). Thermal denaturation temperatures (Tm) were determined
with a Beckman DU 800 spectrophotometer as the maxima of the
[7] A. Pasternak, J. Wengel, Bioog. Med. Chem. Lett. 2011, 21, 752–755.
[8] T. Agarwal, S. Kumar, S. Maiti, Biochimie 2011, 93, 1694–1700.
[9] M. M. Mangos, K.-L. Min, E. Viazovkina, A. Galarneau, M. I. Elzagheid,
M. A. Parniak, M. J. Damha, J. Am. Chem. Soc. 2003, 125, 654–661.
[10] A. MuÇoz-Alarcꢅn, P. Guterstam, C. Romero, M. A. Behlke, K. A. Lennox,
J. Wengel, S. El Andaloussi, ꢆ. Langel, ISRN Pharm. 2012, 2012, 407154.
[11] a) J. B. Bramsen, M. B. Laursen, A. F. Nielsen, T. B. Hansen, C. Bus, N.
Langkjær, B. R. Babu, T. Højland, M. Abramov, A. Van Aerschot, D. Odadz-
ic, R. Smicius, J. Haas, C. Andree, J. Barman, M. Wenska, P. Srivastava, C.
Zhou, D. Honcharenko, S. Hess, E. Muller, G. V. Bobkov, S. N. Mikhailov, E.
ꢂ 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemBioChem 2014, 15, 146 – 156 155

CHEMBIOCHEM
FULL PAPERS
www.chembiochem.org
Fava, T. F. Meyer, J. Chattopadhyaya, M. Zerial, J. W. Engels, P. Herdewijn,
J. Wengel, J. Kjems, Nucleic Acids Res. 2009, 37, 2867–2881; b) D. M.
Kenski, A. J. Cooper, J. J. Li, A. T. Willingham, H. J. Haringsma, T. A.
Young, N. A. Kuklin, J. J. Jones, M. T. Cancilla, D. R. McMasters, M.
Mathur, A. B. Sachs, W. M. Flanagan, Nucleic Acids Res. 2010, 38, 660–
671; c) M. B. Laursen, M. M. Pakula, S. Gao, K. Fluiter, O. R. Mook, F. Baas,
N. Langkjær, S. L. Wengel, J. Wengel, J. Kjems, J. B. Bramsen, Mol. BioSyst.
2010, 6, 862–870; d) D. Werk, J. Wengel, S. L. Wengel, H.-P. Grunert, H.
Zeichhardt, J. Kurreck, FEBS Lett. 2010, 584, 591–598; e) N. Vaish, F.
Chen, S. Seth, K. Fosnaugh, Y. Liu, R. Adami, T. Brown, Y. Chen, P. Harvie,
R. Johns, G. Severson, B. Granger, P. Charmley, M. Houston, M. V. Tem-
plin, B. Polisky, Nucleic Acids Res. 2011, 39, 1823–1832; f) J. B. Bramsen,
M. M. Pakula, T. B. Hansen, C. Bus, N. Langkjær, D. Odadzic, R. Smicius,
S. L. Wengel, J. Chattopadhyaya, J. W. Engels, P. Herdewijn, J. Wengel, J.
Kjems, Nucleic Acids Res. 2010, 38, 5761–5773; g) N. M. Snead, J. R. Esca-
milla-Powers, J. J. Rossi, A. P. McCaffrey, Mol. Ther. Nucleic Acids 2013, 2,
e103.
Chem. Eur. J. 2008, 14, 9968–9980; d) U. V. Schneider, I. Geci, N. Johnk,
N. D. Mikkelsen, E. B. Pedersen, G. Lisby, PLoS One 2011, 6, e20565.
[18] K. K. Karlsen, A. Okholm, J. Kjems, J. Wengel, Bioorg. Med. Chem. 2013,
21, 6186–6190.
[19] T. L. Netzel, M. Zhao, K. Nafisi, J. Headrick, M. S. Sigman, B. E. Eaton, J.
Am. Chem. Soc. 1995, 117, 9119–9128.
[20] a) N. Amann, E. Pandurski, T. Fiebig, H.-A. Wagenknecht, Angew. Chem.
2002, 114, 3084–3087; Angew. Chem. Int. Ed. 2002, 41, 2978–2980;
b) N. Amann, H.-A. Wagenknecht, Synlett 2002, 687–691; c) E. Mayer, L.
Valis, R. Huber, N. Amann, H.-A. Wagenknecht, Synthesis 2003, 2335–
2340.
[21] P. Kaden, E. Mayer-Enthart, A. Trifonov, T. Fiebig, H.-A. Wagenknecht,
Angew. Chem. 2005, 117, 1662–1666; Angew. Chem. Int. Ed. 2005, 44,
1636–1639.
[22] a) E. Mayer-Enthart, H.-A. Wagenknecht, Angew. Chem. 2006, 118, 3451–
3453; Angew. Chem. Int. Ed. 2006, 45, 3372–3375; b) E. Mayer-Enthart,
C. Wanger, J. Barbaric, H.-A. Wagenknecht, Tetrahedron 2007, 63, 3434–
3439.
[23] N. Amann, E. Pandurski, T. Fiebig, H.-A. Wagenknecht, Chem. Eur. J.
2002, 8, 4877–4883.
[24] I. J. Lee, M. Park, T. Joo, B. H. Kim, Mol. BioSyst. 2012, 8, 486–490.
[25] A. S. Wabha, A. Esmaeili, M. J. Damha, R. H. E. Hudson, Nucleic Acids Res.
2010, 38, 1048–1056.
[26] Y.-C. Shih, T.-C. Chien, Tetrahedron 2011, 67, 3915–3923.
[27] A. Rajendran, S. Nakano, N. Sugimoto, Chem. Commun. 2010, 46, 1299–
1301.
[12] a) A. Pasternak, F. J. Hernandez, L. M. Rasmussen, B. Vester, J. Wengel,
Nucleic Acids Res. 2011, 39, 1155–1164; b) T. B. Jensen, J. R. Henriksen,
B. E. Rasmussen, L. M. Rasmussen, T. L. Andresen, J. Wengel, A. Paster-
nak, Bioorg. Med. Chem. 2011, 19, 4739–4745.
[13] K. K. Karlsen, T. B. Jensen, J. Wengel, J. Org. Chem. 2009, 74, 8838–8841.
[14] V. L. Malinovskii, D. Wenger, R. Hꢇner, Chem. Soc. Rev. 2010, 39, 410–
422.
[15] M. E. Østergaard, P. J. Hrdlicka, Chem. Soc. Rev. 2011, 40, 5771–5788.
[16] a) U. B. Christensen, E. B. Pedersen, Nucleic Acids Res. 2002, 30, 4918–
4925; b) U. B. Christensen, E. B. Pedersen, Helv. Chim. Acta 2003, 86,
2090–2097; c) A. Pasternak, E. Kierzek, K. Pasternak, A. Fratczak, D. H.
Turner, R. Kierzek, Biochemistry 2008, 47, 1249–1258.
[28] a) I. J. Lee, J. W. Yi, B. H. Kim, Chem. Commun. 2009, 5383–5385; b) I. J.
Lee, B. H. Kim, Chem. Commun. 2012, 48, 2074–2076.
[29] M. J. Cavaluzzi, P. N. Borer, Nucleic Acids Res. 2004, 32, e13.
[17] a) V. V. Filichev, E. B. Pedersen, J. Am. Chem. Soc. 2005, 127, 14849–
14858; b) V. V. Filichev, I. V. Astakhova, A. D. Malakhov, V. A. Korshun,
E. B. Pedersen, Nucleic Acids Symp. Ser. 2008, 52, 347–348; c) V. V. Fili-
chev, I. V. Astakhova, A. D. Malakhov, V. A. Korshun, E. B. Pedersen,
Received: September 3, 2013
Published online on November 25, 2013
ꢂ 2014 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
ChemBioChem 2014, 15, 146 – 156 156