C2′-pyrene-functionalized triazole-linked DNA: Universal DNA/RNA hybridization probes

Article abstract of DOI:10.1021/jo201845z

Development of universal hybridization probes, that is, oligonucleotides displaying identical affinity toward matched and mismatched DNA/RNA targets, has been a longstanding goal due to potential applications as degenerate PCR primers and microarray probes. The classic approach toward this end has been the use of "universal bases" that either are based on hydrogen-bonding purine derivatives or aromatic base analogues without hydrogen-bonding capabilities. However, development of probes that result in truly universal hybridization without compromising duplex thermostability has proven challenging. Here we have used the "click reaction" to synthesize four C2′-pyrene-functionalized triazole-linked 2′-deoxyuridine phosphoramidites. We demonstrate that oligodeoxyribonucleotides modified with the corresponding monomers display (a) minimally decreased thermal affinity toward DNA/RNA complements relative to reference strands, (b) highly robust universal hybridization characteristics (average differences in thermal denaturation temperatures of matched vs mismatched duplexes involving monomer W are <1.7 °C), and (c) exceptional affinity toward DNA targets containing abasic sites opposite of the modification site (ΔT_m up to +25 °C). The latter observation, along with results from absorption and fluorescence spectroscopy, suggests that the pyrene moiety is intercalating into the duplex whereby the opposing nucleotide is pushed into an extrahelical position. These properties render C2′-pyrene-functionalized triazole-linked DNA as promising universal hybridization probes for applications in nucleic acid chemistry and biotechnology.

Full text of DOI:10.1021/jo201845z

Featured Article
pubs.acs.org/joc
C2′-Pyrene-Functionalized Triazole-Linked DNA: Universal DNA/RNA
Hybridization Probes
Sujay P. Sau and Patrick J. Hrdlicka*
Department of Chemistry, University of Idaho, Moscow, Idaho 83844, United States
S
* Supporting Information
ABSTRACT: Development of universal hybridization probes, that
is, oligonucleotides displaying identical affinity toward matched
and mismatched DNA/RNA targets, has been a longstanding goal
due to potential applications as degenerate PCR primers and
microarray probes. The classic approach toward this end has been
the use of “universal bases” that either are based on hydrogen-
bonding purine derivatives or aromatic base analogues without
hydrogen-bonding capabilities. However, development of probes
that result in truly universal hybridization without compromising
duplex thermostability has proven challenging. Here we have used the “click reaction” to synthesize four C2′-pyrene-
functionalized triazole-linked 2′-deoxyuridine phosphoramidites. We demonstrate that oligodeoxyribonucleotides modified with
the corresponding monomers display (a) minimally decreased thermal affinity toward DNA/RNA complements relative to
reference strands, (b) highly robust universal hybridization characteristics (average differences in thermal denaturation
temperatures of matched vs mismatched duplexes involving monomer W are <1.7 °C), and (c) exceptional affinity toward DNA
targets containing abasic sites opposite of the modification site (ΔTm up to +25 °C). The latter observation, along with results
from absorption and fluorescence spectroscopy, suggests that the pyrene moiety is intercalating into the duplex whereby the
opposing nucleotide is pushed into an extrahelical position. These properties render C2′-pyrene-functionalized triazole-linked
DNA as promising universal hybridization probes for applications in nucleic acid chemistry and biotechnology.
INTRODUCTION
■
The CuI-catalyzed [3 + 2] azide−alkyne cycloaddition
(CuAAC) reaction, which results in the formation of 1,4-
disubstituted 1,2,3-triazoles,1,2 has been extensively utilized for
the synthesis of modified nucleosides, nucleotides, and
oligonucleotides.3,4 The remarkable progress over the past
five years has paved the way for empowering applications in
nucleic acid chemistry5 such as postsynthetic labeling of
oligonucleotides with reporter groups,6 controlled metallization
of oligonucleotides,7 ligation of oligonucleotides,8,9 and
generation of oligonucleotides with artificial backbone10 and
nucleobase11,12 motifs.
Our interest in (a) employing the CuAAC reaction within
oligonucleotide chemistry,13 (b) studying pyrene-functionalized
Figure 1. Structures of C2′-pyrene-functionalized triazole-linked 2′-
deoxyuridine monomers and other monomers studied herein.
oligonucleotide probes for potential diagnostic applications,13−17
and (c) developing oligonucleotides modified with 2′-intercala-
were specifically selected to study the influence of the linker
tor-functionalized nucleotide monomers for DNA-targeting
between the pyrene and triazole moieties on hybridization
applications18−21 prompted us to explore oligodeoxyribonucleo-
properties of correspondingly modified ONs, while non-
tides (ONs) that are modified with C2′-pyrene-functionalized
functionalized monomer V provides insight into the relative
triazole-linked 2′-deoxyuridine monomers W−Z (Figure 1). We
roles of pyrene and triazole moieties. Recent reports have
surmised that the corresponding phosphoramidites would be
described pre-25,26 and postsynthetic6e,27 uses of the CuAAC
readily available via CuAAC reactions using simple reagents and
starting materials, and that the pyrene moieties of monomer
reaction for 2′-functionalization of nucleotides. To the best
of our knowledge, the present work is the first example of
W−Z intercalate into duplex cores as observed with O2′-
intercalator-functionalized RNA,21−23 N2′-intercalator-func-
tionalized 2′-N-methyl-2′-amino-DNA,21,24 and N2′-intercala-
Received: September 7, 2011
Published: November 16, 2011
tor-functionalized 2′-amino-α-L-LNA.18−20 Monomers W−Z
© 2011 American Chemical Society
5
dx.doi.org/10.1021/jo201845z|J. Org. Chem. 2012, 77, 5−16

The Journal of Organic Chemistry
Featured Article
Scheme 1. Synthesis and Structures of Terminal Alkynes
Scheme 2. Synthesis of C2′-Pyrene-Functionalized Triazole-Linked Uridine Phosphoramidites 3V−3Z
ONs modified with monomers where pyrene-functionalized
1,2,3-triazolyl moieties are directly attached to the 2′-position of
nucleosides.
of the resultant homopropargyl alcohol using trifluoroboron
etherate and triethylsilane, afforded Ay in 31% yield.
Room temperature CuAAC reactions between 1 and
terminal alkynes Av−Az provided the corresponding triazoles
2V−2Z in robust yields (60−83%), except when using 1-
ethynylpyrene Aw, which required heating (75 °C) to afford
nucleoside 2W in 35% yield (Scheme 2). Nucleosides 2V−2Z
were subsequently converted into phosphoramidites 3V−3Z
(51−67% yield) using 2-cyanoethyl-N,N,N′,N′-tetraisopropyl-
phosphordiamidite (PN2 reagent) and 1H-tetrazole as an
activator. Alternative phosphitylation conditions were not
investigated as the described route provided sufficient
quantities of 3V−3Z for further analysis.
ON Synthesis and Experimental Design. Phosphorami-
dites 3V−3Z were incorporated into ONs via machine-assisted
solid-phase DNA synthesis (0.2 μmol scale) using the following
conditions (activator, coupling time, stepwise coupling yield):
3V (4,5-dicyanoimidazole, 15 min, ∼80%), 3W (5-(ethylthio)-
1H-tetrazole, 30 min, ∼90%), 3X (5-(bis-3,5-trifluoromethyl-
phenyl)-1H-tetrazole [Activator 42]; 30 min, ∼80%), 3Y and
3Z (4,5-dicyanoimidazole, 30 min, ∼90%). Acceptable but non-
optimized conditions (≥80% coupling yield) were identified
through progressive screening of activators (4,5-dicyanoimida-
zole → 5-(ethylthio)-1H-tetrazole → Activator 42). After
workup and HPLC purification, the composition and purity of
all modified ONs was verified by MALDI TOF MS analysis
(Table S1 in the Supporting Information) and ion-pair reverse-
phase HPLC, respectively.
Here we demonstrate that ONs modified with C2′-pyrene-
functionalized triazole-linked monomers are robust universal
DNA/RNA hybridization probes; that is, they display virtually
identical DNA/RNA target affinity regardless of the nucleotide
opposite of the modification site. Development of universal
hybridization probes has been a longstanding goal due to their
potential as degenerate PCR primers and microarray probes
when the identity of one or more nucleotides in a target
sequence is unknown.28−31 The classic approach toward this
end has been the use of ONs containing “universal bases”,32
which fall into two categories: (a) aromatic base analogues
without hydrogen-bonding capabilities such as 3-nitropyrrole,29
5-nitroindole,33 isocarbostyril,34 or pyrene;13,35−38 and (b)
hydrogen-bonding universal bases based on inosine39−41 or
other purine moieties.42,43 However, development of truly
“universal” hybridization probes that do not compromise
duplex thermostability has proven challenging.44
RESULTS AND DISCUSSION
■
Phosphoramidite Synthesis. 5′-O-Dimethoxytrityl-2′-
azido-2′-deoxyuridine 145 was identified as a suitable starting
material for the synthesis of the target compounds. 2,2,2-
Trifluoro-N-(prop-2-ynyl)acetamide Av,46 1-ethynylpyrene
Aw,47 and N-(prop-2-ynyl)pyrene-1-carboxamide Az48 were
prepared as previously described, while 1-(pyren-1-yl)-prop-2-
yn-1-one Ax and 4-(pyren-1-yl)-but-1-yne Ay were obtained
via novel routes (Scheme 1). Thus, nucleophilic addition of
Me3SiCCMgBr (generated in situ from trimethylsilylacety-
lene and MeMgBr in THF) to pyrene-1-carboxaldehyde
followed by desilylation using potassium carbonate provided
Ax′ in 58% yield. A subsequent Jones oxidation afforded Ax in
75% yield. Similarly, nucleophilic addition of HCCCH2ZnBr
(generated in situ from propargyl bromide and activated zinc in
THF) to pyrene-1-carboxaldehyde, followed by deoxygenation
The hybridization characteristics of ONs modified with
W−Z monomers were examined in 13-mer sequence contexts
that have previously been used to study base-discriminating
fluorescent ONs.13,15,48 Nucleotides flanking the W−Z
monomers were systematically varied to explore the influence
of sequence context on hybridization characteristics (Table 1).
The thermostability of duplexes was evaluated by determining
their thermal denaturation temperature (Tm) in a medium salt
buffer ([Na+] = 110 mM, pH 7.0). Changes in Tm values of
modified duplexes are discussed relative to Tm values of
6
dx.doi.org/10.1021/jo201845z|J. Org. Chem. 2012, 77, 5−16

The Journal of Organic Chemistry
Featured Article
a
Table 1. Tm Values of Duplexes between Centrally Modified ONs and Complementary or Centrally Mismatched DNA Targets
Tm (ΔTm) [°C]
mismatch ΔTm [°C]
ON
sequence
B=
A
C
G
T
avg. mismatch ΔTm seq. [°C] avg. mismatch ΔTm series [°C]
1
2
3
4
5′-CGCAA ATA AACGC
5′-CGCAA CTC AACGC
5′-CGCAA GTG AACGC
5′-CGCAA TTT AACGC
48.5
55.5
55.5
48.5
−10.0
−13.5
−13.0
−11.0
−5.0
−9.5
−9.5
−9.0
−9.0
−9.0
−10.0
−11.0
−8.0 ± 2.6
−10.7 ± 2.5
−10.8 ± 1.9
−10.3 ± 1.2
−10.0 ± 2.2
+0.8 ± 1.9
−0.5 ± 2.2
−1.1 ± 2.1
−4.1 ± 1.6
5
6
7
8
5′-CGCAA AWA AACGC
5′-CGCAA CWC AACGC
5′-CGCAA GWG AACGC
5′-CGCAA TWT AACGC
48.0 (−0.5)
53.5 (−2.0)
51.5 (−4.0)
47.0 (−1.5)
+1.0
+0.5
+1.0
+2.5
+1.5
+2.0
−4.5
−0.5
+1.5
+2.5
0.0
+1.3 ± 0.3
+1.7 ± 1.0
−1.2 ± 2.9
+1.3 ± 1.6
+2.0
9
10
11
12
5′-CGCAA AXA AACGC
5′-CGCAA CXC AACGC
5′-CGCAA GXG AACGC
5′-CGCAA TXT AACGC
46.5 (−2.0)
52.0 (−3.5)
52.5 (−3.0)
44.5 (−4.0)
+1.0
−1.5
+0.5
+1.0
+0.5
0.0
+1.0
−0.5
−0.5
0.0
+0.8 ± 0.3
−0.7 ± 0.8
−2.3 ± 4.1
0.0 ± 1.0
−7.0
−1.0
13
14
15
16
5′-CGCAA AYA AACGC
5′-CGCAA CYC AACGC
5′-CGCAA GYG AACGC
5′-CGCAA TYT AACGC
49.5 (+1.0)
50.5 (−5.0)
53.0 (−2.5)
44.5 (−4.0)
+1.5
−5.0
+1.5
−2.0
0.0
−1.0
−3.5
−2.0
+1.0
−2.5
+0.5
−1.5
+0.8 ± 0.8
−2.8 ± 2.0
−0.5 ± 2.6
−1.8 ± 0.3
17
18
19
20
5′-CGCAA AZA AACGC
5′-CGCAA CZC AACGC
5′-CGCAA GZG AACGC
5′-CGCAA TZT AACGC
47.0 (−1.5)
51.5 (−4.0)
52.0 (−3.5)
45.5 (−3.0)
−5.5
−6.5
−2.0
−4.5
−2.5
−1.0
−6.0
−5.0
−4.0
−4.0
−4.0
−4.0
−4.0 ± 1.5
−3.8 ± 2.8
−4.0 ± 2.0
−4.5 ± 0.5
a
Tm values determined as maximum of the first derivative of denaturation curves (A260 vs T) recorded in thermal denaturation buffer ([Na+] = 110
mM, [Cl−] = 100 mM, pH 7.0 (NaH2PO4/Na2HPO4)) using 1.0 μM of each strand. Tm values are averages of at least two measurements within
1.0 °C. ΔTm = change in Tm relative to unmodified reference duplex. Mismatch ΔTm = change in Tm relative to fully matched duplex (B = A). Avg.
mismatch ΔTm seq = average of all three mismatch ΔTm values for a given probe. Avg. mismatch ΔTm series = average of all 12 mismatch ΔTm
values of all four studied probe contexts within a monomer series; ± denotes standard deviation. DNA targets: 3′-GCGTT TBT TTGCG (for ON1/
ON5/ON9/ON13/ON17), 3′-GCGTT GBG TTGCG (for ON2/ON6/ON10/ON14/ON18), 3′-GCGTT CBC TTGCG (for ON3/ON7/
ON11/ON15/ON19), and 3′-GCGTT ABA TTGCG (for ON4/ON8/ON12/ON16/ON20). For structures of monomers W−Z, see Figure 1.
unmodified reference duplexes (ΔTm). The exchange of
thymine (reference ONs) with uracil moieties (modified
ONs) results in a decrease of 0.5 °C per incorporation49 by
itself but is not considered further herein.
duplexes exhibit minimal changes in Tm values relative to
matched duplexes (mismatch ΔTm values between −1.5 and
+2.5 °C, Table 1). ONs with a GBG context that are hybridized
to dG-mismatched targets are the exception hereto (see
mismatch ΔTm values for ON7 and ON11, Table 1). Thus,
the average mismatch ΔTm values across the four studied
sequence contexts are +0.8 and −0.5 °C for ONs modified with
monomer W and X, respectively. In contrast, unmodified
reference strands ON1−ON4 display the expected mismatch
discrimination profile including (a) formation of substantially
Thermal Denaturation Studies. Thermal denaturation
curves of DNA duplexes modified with C2′-pyrene-functional-
ized triazole-linked 2′-deoxyuridine monomers W−Z display
similar sigmoidal monophasic transitions as unmodified
reference duplexes (Figure S1). ONs that are centrally modified
with a single W−Z monomer generally display moderately
decreased thermal affinity toward complementary DNA (ΔTm
for ON5−ON20 between −5.0 and +1.0 °C, Table 1). Less
pronounced destabilization is observed for (a) ONs modified
with monomer W where the pyrene is directly linked to the
triazole moiety, and (b) ONs with a central ABA-context
(ON5/ON9/ON13/ON17).
Control studies using 9-mer ONs revealed that monomer V,
which lacks a pyrene moiety, induces larger decreases in duplex
thermostability than pyrene-functionalized monomer Z (differ-
ence in Tm values = 2.5−4.5 °C, Table S2). This indicates that
the C2′-triazole moiety is the primary destabilizing structural
element of the W−Z monomers.
destabilized mismatched duplexes (average mismatch ΔTm
=
−10.0 °C, Table 1) and (b) more efficient discrimination of
pyrimidine−pyrimidine mismatches than of pyrimidine−purine
mismatches. ONs modified with monomer Y, where the pyrene
and triazole moieties are separated by a flexible two-carbon
linker (ON13−ON16), also display universal hybridization
characteristics albeit with slightly greater sequence- and
mismatch-dependent variation than observed for ON5−
ON12 (average mismatch ΔTm = −1.1 °C, Table 1). ONs
modified with monomer Z, which has the longest linker studied
herein, do not display universal hybridization characteristics.
However, markedly reduced discrimination of mismatched
targets is still observed (compare mismatch ΔTm values for
ON17−ON20 and ON1-ON4, Table 1).
The Watson−Crick specificity of singly modified ON5−
ON20 was studied using DNA targets with mismatched
nucleotides opposite of the modification site. Interestingly,
ON5−ON12 (monomers W/X) display extremely robust
universal hybridization characteristics; that is, mismatched
We have previously studied ONs modified with C5-pyrene-
functionalized triazole-linked 2′-deoxyuridine monomers in
identical sequence contexts and found them to display similar
7
dx.doi.org/10.1021/jo201845z|J. Org. Chem. 2012, 77, 5−16

The Journal of Organic Chemistry
Featured Article
a
Table 2. Tm Values of Duplexes between ON21−ON33 and Complementary or Centrally Mismatched DNA Targets
DNA: 5′-CGCAA ABA AACGC
Tm (ΔTm) [°C]
mismatch ΔTm [°C]
ON
sequence
B=
T
A
C
G
21
3′-GCGTT TAT TTGCG
48.5
−10.0
−10.0
−5.5
22
23
24
3′-GCGTT TAW TTGCG
3′-GCGTT WAT TTGCG
3′-GCGTT WAW TTGCG
46.5 (−2.0)
41.0 (−7.5)
35.0 (−13.5)
−17.0
−11.5
nt
−9.0
−10.0
−3.5
−12.0
−5.5
nt
25
26
27
3′-GCGTT TAX TTGCG
3′-GCGTT XAT TTGCG
3′-GCGTT XAX TTGCG
44.0 (−4.5)
42.0 (−6.5)
36.5 (−12.0)
−9.0
−10.5
−2.5
−7.5
−11.5
−4.5
−5.0
−6.0
+0.5
28
29
30
3′-GCGTT TAY TTGCG
3′-GCGTT YAT TTGCG
3′-GCGTT YAY TTGCG
40.0 (−8.5)
42.5 (−6.0)
34.5 (−14.0)
−7.0
−6.0
−4.5
−7.0
−2.5
−3.5
−5.0
−7.0
−4.5
31
32
33
3′-GCGTT TAZ TTGCG
3′-GCGTT ZAT TTGCG
3′-GCGTT ZAZ TTGCG
42.5 (−6.0)
44.0 (−4.5)
39.5 (−9.0)
−7.0
−11.5
−2.5
−8.5
−12.0
−0.5
−7.5
−9.0
−3.5
a
Conditions and definitions as described in footnote of Table 1; nt = no transition.
mismatch ΔTm values as ONs modified with monomer W/X/
Y; however, significantly greater duplex destabilization was
observed (average ΔTm ∼ −7.5 °C).13 In contrast, ONs
modified with the related 2′-O-(pyren-1-yl)methyluridine or 2′-
N-(pyren-1-ylmethyl)-2′-N-methylaminouridine monomers dis-
play very high thermal affinity toward complementary DNA
and do not display universal hybridization characteristics.21
These observations suggest that pyrene-functionalized triazole
moieties are structural units that promote universal hybrid-
ization characteristics, and that their specific attachment point
on the nucleotide influences duplex thermostability.
Next, ON22−ON33 were prepared to study how incorpo-
ration of W−Z monomers as next-nearest neighbors influences
duplex thermostability and if the presence of W−Z monomers
influences the discriminatory ability of the neighboring
nucleoside for its Watson−Crick complement (Table 2).50
Singly modified ONs with TBA and ABT contexts display
lower thermal affinity toward complementary DNA than ONs
with symmetric ABA or TBT contexts (e.g., compare ΔTm
values for ON5, ON8, ON22, and ON23, Table 2). This
underscores the general point that new nucleotide monomers
must be studied in many different sequence contexts before a
full understanding of hybridization effects is reached.
Incorporation of a second X−Z monomer results in
approximately additive decreases in duplex thermostability,
while greater-than-additive decreases are observed with
monomer W (e.g., compare ΔTm values for ON22/ON23/
ON24, Table 2).
values for ON33 relative to ON21, Table 2), although these
trends cannot be categorized as universal hybridization. We
speculate that the poor mismatch specificity is caused by the
dynamic local duplex structure that arises as a consequence of
the low duplex thermostability.
In summary, the data demonstrate that ONs modified with
C2′-pyrene-functionalized triazole-linked 2′-deoxyuridine
monomers display universal hybridization characteristics with
DNA targets that have mismatched nucleotides opposite of the
modification site (compare mismatch ΔTm values in Tables 1
and 2 but only have limited influence on the Watson−Crick
specificity of neighboring base pairs).
As a first step toward rationalizing whether intercalation of
the pyrene/triazole moieties of monomers W−Z governs the
observed universal hybridization characteristics, we hybridized
ON4/ON8/ON12/ON16/ON20 (TBT context) to DNA
targets containing a THF-type abasic site monomer Φ51
opposite of monomers W−Z (for structure of monomer Φ, see
Figure 1). As expected, the duplex between reference strand
ON4 and the abasic target strand is greatly destabilized relative
to the matched duplex due to perturbation of the base stack
(abasic ΔTm = −20.0 °C, Table 3). In contrast, ONs modified
with monomers W−Z result in the formation of remarkably
thermostable duplexes with abasic target strands (abasic ΔTm
between −3.5 and +4.5 °C, Table 3). The observed trend in
abasic ΔTm values (W > X > Y > Z) demonstrates that
monomers with progressively longer linkers between the
pyrene and triazole moieties result in progressively less
pronounced stabilization of abasic sites.
Stabilization of abasic sites has previously been observed for
monomers with extended aromatic units which occupy the void
formed by an abasic site and re-establish π−π stacking at the
lesion site.18,35,52−54 Full restoration of duplex thermostability,
however, is rarely observed. These observations indicate that
the pyrene and/or triazole moieties of monomers W−Z are
intercalating into the duplex core and thereby disrupt
interactions between mismatched base pairs, leading to a lack
The presence of a single W−Z monomer has, with few
exceptions (ON22/ON29), only a minor effect on the
discriminatory ability of neighboring base pairs (e.g., compare
mismatch ΔTm values for ON31/ON32 relative to ON21,
Table 2). This reduces the risk of nonspecific target binding
and suggests that the pyrene-functionalized triazole units of
W−Z monomers do not interact strongly with neighboring
base pairs. Doubly modified ONs display poor discrimination
of DNA targets with mismatched nucleotides positioned
between the modification sites (e.g., compare mismatch ΔTm
8
dx.doi.org/10.1021/jo201845z|J. Org. Chem. 2012, 77, 5−16

The Journal of Organic Chemistry
Featured Article
Table 3. Tm Values of Duplexes between Centrally Modified ONs (TBT Context) and Complementary DNA or Targets
a
Containing a Central Abasic Site
3′-GCGTT ATA TTGCG
Tm [°C]
3′-GCGTT AΦA TTGCG
abasic ΔTm [°C]
ON
sequence
4
8
5′-CGCAA TTT AACGC
5′-CGCAA TWT AACGC
5′-CGCAA TXT AACGC
5′-CGCAA TYT AACGC
5′-CGCAA TZT AACGC
48.5
47.0
44.5
44.5
45.5
−20.0
+4.5
+2.0
+0.5
−3.5
12
16
20
a
Conditions as described in footnote of Table 1. Abasic ΔTm = change in Tm relative to fully matched duplex; Φ = abasic monomer (for structure,
see Figure 1).
Figure 2. Representative absorption spectra of single-stranded ON8/ON12/ON16/ON20 (a−d) and their duplexes with matched (M) and
centrally mismatched (MM) DNA targets: 3′-GCGTT ABA TTGCG. Nucleotide opposite of modification is mentioned in parentheses. Spectra
were recorded in thermal denaturation buffer at T = 20 °C using 1.0 μM concentration of each strand. Note that different x-axes are used. “O”
denotes THF-type abasic site monomer Φ (Figure 1).
of thermal preference for a particular nucleotide opposite of the
modification site.
suggests electronic coupling between the pyrene and triazole
moieties. Single-stranded ON13−ON16 (monomer Y) and
ON17−ON20 (monomer Z), on the other hand, have
structured absorption spectra with two maxima in the “normal”
region (i.e., λmax ∼ 333/348 and ∼332/346 nm, respectively,
Figure 2). Hybridization of ON13−ON20 with complemen-
tary, mismatched, or abasic DNA target strands results in subtle
bathochromic shifts (Δλmax between +1 and +3 nm, Figure 2
and Table S3). Thus, the absorption data are consistent with
the hypothesis that the pyrene moieties of monomers W−Z
intercalate into the duplex core upon hybridization with DNA
targets.
Next, steady-state fluorescence emission spectra and
fluorescence emission quantum yields were determined for
ON5−ON20 in the absence or presence of complementary or
centrally mismatched DNA targets (Figure 3 and Table 4).
Monomer W. Single-stranded ON5−ON8 display two
structured emission peaks at λem ∼ 390 and 405 nm (Figure 3).
The single-stranded probe with a central AWA context
(ON5) has higher fluorescence quantum yield than SSPs in
other contexts (Φ F = 0.27 vs 0.07/0.05/0.05, Table 4; see
also Figure S2). This is in agreement with previous
observations that adenine is the weakest quencher of pyrene
fluorescence (quenching trend: G > C > T > A).15,56,57 The
Optical Spectroscopy Studies. UV−vis absorption spec-
tra of ONs modified with monomers W−Z were recorded in
the absence or presence of complementary or centrally
mismatched DNA targets in order to gain additional insights
into the mechanism that governs the observed universal
hybridization characteristics (Figure 2); hybridization-induced
intercalation of pyrene moieties is known to induce subtle
bathochromic shifts.55
Single-stranded ON5−ON8 (monomer W) display a single
unstructured maximum in the pyrene region (λ max ∼ 351 nm,
Figure 2), while duplexes with complementary, mismatched, or
abasic DNA targets display two resolved maxima at ∼351 and
∼365 nm. The lack of defined peaks for the single-stranded
probes (SSPs) precludes analysis of bathochromic shifts. Single-
stranded ON9−ON12 (monomer X) display two broad and
virtually equally intense peaks, which renders exact determi-
nation of absorption maxima difficult (λ max ∼ 385 and ∼415
nm, Figure 2). Hybridization with complementary DNA results
in subtle bathochromic shifts, while more pronounced shifts are
observed upon hybridization with mismatched or abasic DNA.
The pyrene maxima of ON5−ON12 are red-shifted relative to
those of unconjugated pyrene chromophores,13,19,21 which
9
dx.doi.org/10.1021/jo201845z|J. Org. Chem. 2012, 77, 5−16

The Journal of Organic Chemistry
Featured Article
Figure 3. Steady-state fluorescence emission spectra of single-stranded ON8/ON12/ON16/ON20 (TTT context) and duplexes with matched (M)
or centrally mismatched (MM) DNA. Recorded in thermal denaturation buffer at T = 20 °C using 1.0 μM of each strand and λex = 350 nm
(monomers W, Y, and Z) or λex = 400 nm (monomer X). DNA targets 3′-GCGTT ABA TTGCG. Nucleotide opposite of modification is mentioned
in parentheses. Note that different axes are used. “O” denotes THF-type abasic site monomer Φ (Figure 1).
Table 4. Relative Fluorescence Emission Quantum Yield (Φ F) of ON5−ON20 in the Absence (SSP) or Presence of Matched
a
(M) or Centrally Mismatched (MM) DNA Targets
ΦF
ON
sequence
SSP
+M (A)
+MM (C)
+MM (G)
+MM (T)
5
6
7
8
5′-CGCAA AWA AACGC
5′-CGCAA CWC AACGC
5′-CGCAA GWG AACGC
5′-CGCAA TWT AACGC
0.27
0.07
0.05
0.05
0.08
0.02
0.03
0.07
0.09
0.02
0.01
0.06
0.06
0.02
0.02
0.06
0.08
0.01
0.01
0.04
9
10
11
12
5′-CGCAA AXA AACGC
5′-CGCAA CXC AACGC
5′-CGCAA GXG AACGC
5′-CGCAA TXT AACGC
0.02
0.02
0.25
<0.01
<0.01
0.16
0.33
<0.01
<0.01
0.35
0.10
<0.01
<0.01
0.04
0.25
<0.01
<0.01
0.32
<0.01
0.04
13
14
15
16
5′-CGCAA AYA AACGC
5′-CGCAA CYC AACGC
5′-CGCAA GYG AACGC
5′-CGCAA TYT AACGC
0.09
0.01
0.03
0.01
0.02
0.02
0.02
<0.01
0.01
0.02
<0.01
0.01
0.03
<0.01
<0.01
<0.01
0.01
<0.01
<0.01
<0.01
17
18
19
20
5′-CGCAA AZA AACGC
5′-CGCAA CZC AACGC
5′-CGCAA GZG AACGC
5′-CGCAA TZT AACGC
0.58
0.24
0.05
0.27
0.52
0.15
0.04
0.57
0.78
0.17
0.02
0.58
0.29
0.15
0.02
0.31
0.79
0.19
0.03
0.52
a
Relative to quantum yield of anthracene in ethanol (0.27). Recorded in thermal denaturation buffer at T = 20 °C using 1.0 μM concentration of
each strand and λex = 350 nm and λem = 360−510 nm (monomers W, Y, and Z) or λex = 400 nm and λem = 425−625 nm (monomer X). For DNA
targets, see footnote of Table 1. Nucleotide opposite of modification is mentioned in parentheses.
spectra of the corresponding duplexes with complementary
DNA have a similar shape and sequence dependency,
underlining that the pyrene moiety is in close contact with
the neighboring nucleobases (Figure 3 and Table 4). The
extensive decreases in fluorescence quantum yield (Table 4
and Figure S3) upon hybridization with matched or
mismatched DNA targets further corroborates this hypoth-
esis. ON8 (TWT context) exhibits considerably smaller
changes, presumably since the fluorophore interacts with the
neighboring and only weakly quenching adenine moieties
upon target binding (Figure 3 and Figure S3).
Monomer X. Fluorescence emission spectra of single-
stranded ON9−ON12 and the corresponding duplexes with
complementary or mismatched DNA targets display broad and
unstructured emission peaks with maxima at λem ∼ 490 nm
(Figure 3). SSPs are strongly quenched with ON11 (GXG
context) displaying the lowest intensity (Φ F < 0.04, Table 4
and Figure S2). Quantum yields are markedly increased upon
10
dx.doi.org/10.1021/jo201845z|J. Org. Chem. 2012, 77, 5−16

The Journal of Organic Chemistry
Featured Article
hybridization of ON9 or ON12 with complementary/
mismatched DNA targets (Table 4 and Figure S3). In contrast,
ON10 or ON11 display hybridization-induced decreases in
fluorescence intensity (Table 4 and Figure S3). One inter-
pretation of these observations is that the conjugated pyrene
moiety of monomer X intercalates into the base stack where it is
quenched by neighboring cytosine and guanine moieties (ON10/
ON11) but not quenched by adenine and thymine moieties
(ON9/ON12). An alternative interpretation is that the pyrene
moiety of monomer X only intercalates with ON10/ON11.
However, the similar influence on duplex thermostability upon
incorporation of monomer X irrespective of sequence context
(compare ΔTm values for ON9−ON12, Table 1) and the
hybridization-induced bathochromic shifts of pyrene absorption
peaks (Figure 2) are in stronger support of the first interpretation.
Monomer Y. The fluorescence emission spectra of ON13−
ON16 and the corresponding duplexes with matched or
mismatched DNA targets display two well-resolved pyrene
peaks at λem ∼ 380 and 400 nm, with an additional shoulder at
λem ∼ 420 nm (Figure 3). Very low quantum yields are observed
(ΦF < 0.03, Table 4), except for the single-stranded ON13 (AYA
context). Hybridization of ON13−ON16 with complementary or
mismatched DNA targets generally results in decreased
fluorescence intensity (Figure S3), which is consistent with an
intercalating binding mode for the pyrene moiety.
ON17, and ON20 are exceptions hereto as lower quantum
yields are observed upon hybridization with dG-mismatched
targets than with other DNA targets; however, this most likely
reflects the fact that guanine is a strong fluorophore quencher.56
Collectively, these observations indicate (a) that the
fluorophore is in a similar electronic environment within the
duplex core regardless of the nucleotide opposite of the
monomer and, therefore, (b) that the opposing nucleotide is
not strongly involved in base pairing and possibly even pushed
into an extrahelical position (Figure 4). Along the lines, it is
Figure 4. Illustration of putative mechanism resulting in universal
hybridization.
interesting to note that placement of pyrene-functionalized C-
glycosides in DNA duplexes opposite of abasic sites, which are
generated via enzyme-mediated extrahelical flipping of the
opposing nucleotide, is known to be stabilizing.58,59
Monomer Z. The fluorescence emission spectra of single-
stranded ON17−ON20 and the corresponding duplexes with
matched or mismatched DNA targets display an unstructured
peak at λem ∼ 410 nm with a weaker shoulder at λem ∼ 390 nm
(Figure 3). The quantum yields of SSPs range from moderate
to high and closely align with the previously discussed
quenching trends of nucleobases (ΦF = 0.05−0.58, Table 4
and Figure S2). Hybridization with matched or mismatched
DNA targets generally results in decreases (CZC/GZG
contexts) or minor increases (AZA/TZT contexts) in quantum
yields and intensity (Table 4 and Figure S3), which resembles
the trends with ON9−ON12.
Perhaps the most important observation toward rationalizing
the universal hybridization properties of ON5−ON20 is that
very similar quantum yields are observed for the four duplexes
between a particular probe and matched/mismatched DNA
targets (e.g., compare ΦF = 0.08/0.09/0.06/0.08 for ON5 vs
matched/mismatched DNA targets, Table 4). ON9, ON12,
Universal HybridizationRNA Targets. A representa-
tive subset of modified ONs (TBT/CBT contexts) was studied
with respect to thermal denaturation, absorption, and
fluorescence properties with complementary/mismatched
RNA targets. The following observations were made: (a)
incorporation of monomer W or X into ONs results in similar
decreases in thermal affinity toward complementary RNA as
toward DNA, while ONs modified with monomers Y or Z have
lower affinity toward RNA (Table 5 and Figure S4); (b) ONs
modified with monomers W or X display robust universal
hybridization characteristics (compare mismatch ΔTm values
for ON6/ON8/ON12 and ON2/ON4, Table 5), while ONs
modified with monomers Y or Z do not; (c) pyrene absorption
spectra of duplexes between modified ONs and complementary
or centrally mismatched RNA targets are very similar to those
of the corresponding DNA duplexes (compare Figure S5 and
Figure 2); hybridization-induced bathochromic shifts with
ON14/ON16/ON18/ON20 (monomer Y/Z) are more subtle
a
Table 5. Tm Values of Duplexes between Centrally Modified ONs and Complementary or Centrally Mismatched RNA Targets
Tm (ΔTm) [°C]
mismatch ΔTm [°C]
ON
sequence
B=
A
C
G
U
2
4
5′-CGCAA CTC AACGC
5′-CGCAA TTT AACGC
51.5
40.5
−15.5
−19.0
−3.0
−3.5
−13.5
−17.0
6
8
5′-CGCAA CWC AACGC
5′-CGCAA TWT AACGC
47.0 (−4.5)
42.0 (+1.5)
+1.0
+3.0
+0.0
+0.5
+0.5
+1.5
12
5′-CGCAA TXT AACGC
38.0 (−2.5)
+1.0
+0.0
+1.0
14
16
5′-CGCAA CYC AACGC
5′-CGCAA TYT AACGC
43.5 (−8.0)
36.0 (−4.5)
−2.0
+1.5
−5.0
−1.0
−4.0
−1.0
18
20
5′-CGCAA CZC AACGC
5′-CGCAA TZT AACGC
44.5 (−7.0)
36.5 (−4.0)
−3.0
−12.0
−8.0
−7.5
−7.0
−13.0
a
Conditions and definitions as described in footnote of Table 1. RNA targets: 3′-GCGUU GBG UUGCG (for ON2/ON6/ON14/ON18) and 3′-
GCGUU ABA UUGCG (for ON4/ON8/ON12/ON16/ON20).
11
dx.doi.org/10.1021/jo201845z|J. Org. Chem. 2012, 77, 5−16

The Journal of Organic Chemistry
Featured Article
CH2Cl2 and MeOH (10 mL, 1:1, v/v) and stirred with K2CO3
(0.50 g, 3.62 mmol) at rt for 2 h. The reaction mixture was then
diluted with CH2Cl2 (10 mL) and successively washed with brine (20
mL) and water (20 mL). The organic phase was dried over Na2SO4
and evaporated to dryness. The resulting crude was purified by silica
gel column chromatography (0−50% EtOAc in petroleum ether, v/v)
to afford Ax′ (0.47 g, 58%) as a white solid material: Rf = 0.3 (25%
EtOAc in petroleum ether, v/v); ESI-HRMS m/z 279.0783 ([M +
Na]+, C19H12O·Na+, calcd 279.0780); 1H NMR (DMSO-d6) δ 8.59 (d,
1H, J = 10.0 Hz, Py), 8.35−8.29 (m, 4H, Py), 8.26 (d, 1H, J = 10.0 Hz,
Py), 8.20−8.16 (m, 2H, Py), 8.09 (t, 1H, J = 7.5 Hz, Py), 6.41−6.39
(d, 1H, ex, J = 5.0 Hz, OH), 6.34−6.31 (dd, 1H, J = 5.0 Hz, 2.5 Hz,
HC(OH)), 3.60 (d, 1H, J = 2.5 Hz, HCC); 13C NMR (DMSO-d6)
δ 135.0, 130.7, 130.5, 130.1, 127.34, 127.31 (Py), 127.28 (Py), 127.25
(Py), 126.2 (Py), 125.3 (Py), 125.2 (Py), 124.65 (Py), 124.59 (Py),
124.1, 123.8, 123.7 (Py), 85.5, 76.6 (HCC), 60.8 (HC(OH)).
1-(Pyren-1-yl)-prop-2-yn-1-one (Ax). The Jones reagent (2.67
M CrO3 in 3 M H2SO4, 1.0 mL, 2.67 mmol) was added to a solution
of alcohol Ax′ (180 mg, 0.67 mmol) in acetone (10 mL), and the
reaction mixture was stirred under an ambient atmosphere at rt for 2 h,
whereupon it was diluted with EtOAc (20 mL), neutralized by
dropwise addition of 6 M NaOH (1.0 mL) under stirring, and
sequentially washed with water (30 mL) and saturated aqueous
NaHCO3 (30 mL). The organic phase was dried over Na2SO4,
evaporated to dryness, and the resulting crude purified by silica gel
column chromatography (0−20% EtOAc in petroleum ether, v/v) to
furnish Ax (130 mg, 75%) as a brightly yellow solid material: Rf = 0.6
(50% EtOAc in petroleum ether, v/v); ESI-HRMS m/z 277.0626
([M + Na]+, C19H10O·Na+, calcd 277.0624); 1H NMR (CDCl3) δ 9.48
(d, 1H, J = 10.0 Hz), 8.94 (d, 1H, J = 8.0 Hz), 8.28−8.23 (m, 3H),
8.19−8.14 (m, 2H), 8.07−8.02 (m, 2H), 3.53 (s, 1H); 13C NMR
(CDCl3) δ 179.3, 135.8, 132.2 (Py), 131.31, 131.25 (Py), 131.14,
131.05 (Py), 130.6, 128.4, 127.35 (Py), 127.29 (Py), 127.1 (Py), 126.8
(Py), 124.97 (Py), 124.96, 124.95, 124.2 (Py), 124.1, 82.6, 80.1 (HC
with RNA targets than with the corresponding DNA targets
(compare Table S4 and Table S3); and (d) hybridization of
modified ONs to RNA targets results in very similar changes in
fluorescence intensity as with DNA targets (compare Figure S6
and Figure S3).
Thus, the results indicate that the universal RNA hybrid-
ization characteristics of ONs modified with monomer W/X
(ON6/ON8/ON12/ON20) are governed by a similar
mechanism as universal DNA hybridization (Figure 4).
CONCLUSION
■
Oligodeoxyribonucleotides modified with C2′-pyrene-functional-
ized triazole-linked 2′-deoxyuridine monomers display highly
robust universal hybridization characteristics without markedly
compromising duplex thermostability (Tables 1 and 5), which
sets them apart from probes based on conventional univer-
sal bases such as 3-nitropyrrole or 5-nitroindole. Thermal
denaturation and optical spectroscopy data suggest the universal
hybridization characteristics to be a consequence of pyrene
intercalation whereby the nucleotide opposite of the monomer is
pushed out (Figure 4). Given the straightforward access to this
monomer class via the CuI-catalyzed [3 + 2] azide−alkyne
cycloaddition reaction (Scheme 2), the stage is set for detailed
structure−property studies for refinement of hybridization
characteristics (e.g., attachment of other aromatic moieties) and
biotechnological exploration of these universal hybridization
probes as degenerate PCR primers and microarray probes.
EXPERIMENTAL SECTION
■
General. Reagents and solvents were obtained from commercial
vendors, of analytical grade, and used without further purification.
Petroleum ether of the distillation range 60−80 °C was used. Solvents
were dried over activated molecular sieves: CH2Cl2 and N,N′-
diisopropylethylamine (4 Å). Water content of anhydrous solvents
was verified using Karl Fischer apparatus. Reactions were conducted
under argon whenever anhydrous solvents were used. Reactions were
monitored by TLC using silica gel coated plates with a fluorescence
indicator (SiO2-60, F-254) and were visualized (a) under UV light
and/or (b) by dipping in 5% concd H2SO4 in absolute ethanol (v/v)
followed by heating. Silica gel column chromatography was performed
with silica gel 60 (particle size 0.040−0.063 mm) using moderate
pressure (pressure ball). Evaporation of solvents was carried out under
reduced pressure at temperatures below 45 °C. After column
chromatography, appropriate fractions were pooled, evaporated, and
dried at high vacuum for at least 12 h to give the obtained products in
high purity (>95%) as ascertained by 1D NMR techniques. Chemical
1
C). We observe distinctly different H NMR signals in the 8.50−8.00
ppm region compared to previous reports on this compound.60
4-(Pyren-1-yl)-but-1-yne (Ay). An oven-dried flask was charged
with pyrene-1-carboxaldehyde (230 mg, 1.00 mmol) and activated zinc
(100 mg, 1.50 mmol) and placed under an argon atmosphere.
Anhydrous THF (5 mL) and propargyl bromide (0.20 mL, 1.79
mmol) were added, and the reaction mixture was stirred at 45 °C for 4
h. Saturated aqueous NH4Cl (1 mL) was added, and the mixture was
extracted with EtOAc (2 × 20 mL). The organic phase was washed
with brine (20 mL) and evaporated to dryness. The resulting crude
was purified by silica gel column chromatography (0−30% EtOAc in
petroleum ether, v/v) to afford a crude white solid material (145 mg),
1
which H NMR suggested to be a ∼9:1 mixture of the desired 1-
(pyren-1-yl)-but-3-yn-1-ol and the corresponding allene. Et3SiH (0.20
mL, 1.25 mmol) and boron trifluoride etherate (0.20 mL, 1.62) were
added to a solution of the crude mixture in CH2Cl2 (5 mL), which
then was stirred at rt for 1 h. The reaction mixture was diluted with
CH2Cl2 (20 mL) and saturated aqueous NaHCO3 (2 mL) and
successively washed with brine (20 mL) and water (20 mL). The
organic phase was dried over Na2SO4, evaporated to dryness under
reduced pressure, and the resulting crude purified by silica gel column
chromatography (0−3% EtOAc in petroleum ether, v/v) to afford Ay
(80 mg, 31%) as a white solid material: Rf = 0.5 (5% EtOAc in
petroleum ether, v/v); ESI-HRMS m/z 277.0973 ([M + Na]+,
C20H14·Na+, calcd 279.0988); 1H NMR (CDCl3) δ 8.27−8.25 (d, 1H,
J = 9.5 Hz, Py), 8.17−8.14 (m, 2H, Py), 8.12−8.10 (m, 2H, Py), 8.01
(ap s, 2H), 8.00−7.96 (t, 1H, J = 8.0 Hz, Py), 7.92−7.90 (d, 1H, J =
7.5 Hz, Py), 3.59 (t, 2H, J = 7.7 Hz, CH2CH2CCH), 2.72 (dt, 2H,
J = 7.7 Hz, 2.5 Hz, CH2CCH), 2.03 (t, 1H, J = 2.5 Hz, HCC);
13C NMR (CDCl3) δ 134.7, 131.6, 131.1, 130.5, 128.9, 127.8 (Py),
1
shifts of H NMR (500 MHz), 13C NMR (125.6 MHz), 31P NMR
(121.5 MHz), and/or 19F NMR (282.2 MHz) signals are reported
relative to deuterated solvent or other internal standards (80%
phosphoric acid for 31P NMR). Exchangeable (ex) protons were
1
detected by disappearance of H NMR signals upon D2O addition.
Assignments of NMR spectra are based on 2D spectra (HSQC,
COSY) and DEPT spectra. Quarternary carbons are not assigned in
13C NMR but verified from DEPT spectra (absence of signals).
MALDI-HRMS spectra of compounds were recorded on a Q-TOF
mass spectrometer using 2,5-dihydroxybenzoic acid (DHB) as a matrix
and polyethylene glycol (PEG 600) as an internal calibration standard.
1-(Pyren-1-yl)-prop-2-yn-1-ol (Ax′). Trimethylsilylacetylene
(1.0 mL, 7.00 mmol) was added to MeMgBr in THF (1M, 4.0 mL,
4.00 mmol) under an argon atmosphere and stirred at rt for 1 h. At
this point, pyrene-1-carboxaldehyde (0.70 g, 3.00 mmol) was added
and the reaction mixture was stirred at rt for another 2 h. Saturated
aqueous NH4Cl (∼1 mL) was added, and the mixture was extracted
with EtOAc (2 × 20 mL). The combined organic phase was dried over
Na2SO4 and evaporated to dryness. The resulting crude [assumed to
be 1-(pyren-1-yl)-3-trimethylsilyl-prop-2-yn-1-ol] was dissolved in
127.7 (Py), 127.5 (Py), 127.1 (Py), 126.1 (Py), 125.31, 125.27 (Py),
125.2, 125.1 (Py), 125.0 (Py), 123.2 (Py), 84.0, 69.6 (HCC), 32.8
(CH2CH2CCH), 21.0 (CH2CCH).
General Click Reaction Protocol for Preparation of 2V−2Z
(Description for ∼6 mmol Scale). 5′-O-Dimethoxytrityl-2′-azido-2′-
deoxyuridine 145 and the appropriate alkyne A were added to a
12
dx.doi.org/10.1021/jo201845z|J. Org. Chem. 2012, 77, 5−16

The Journal of Organic Chemistry
Featured Article
mixture of THF/t-BuOH/H2O (3:1:1, v/v/v) along with sodium
ascorbate and CuSO4·5H2O (reagent quantities, and solvent volumes
are specified below). The reaction mixture was stirred under a nitrogen
atmosphere until analytical TLC indicated full conversion (reaction
times and temperatures specified below), whereupon it was diluted
with EtOAc (10 mL). The organic phase was successively washed with
saturated aqueous NaHCO3 (20 mL) and brine (20 mL), dried over
anhydrous Na2SO4, and evaporated to dryness. The resulting crude
was purified by silica column chromatography (eluent specified below)
to afford the corresponding nucleoside 2 (yield specified below).
5′-O-(4,4′-Dimethoxytrityl)-2′-C-[4-(2,2,2-trifluoroacetami-
domethyl)-1H-1,2,3-triazol-1-yl]-2′-deoxyuridine (2V). Nucleo-
side 1 (0.40 g, 0.70 mmol), 2,2,2-trifluoro-N-(prop-2-ynyl)acetamide
Av46 (105 mg, 0.70 mmol), sodium ascorbate (70 mg, 0.35 mmol),
CuSO4·5H2O (5 mg, 0.02 mmol), and THF/t-BuOH/H2O (5 mL)
were mixed, reacted (14 h at rt), worked up, and purified (50−100%
EtOAc in petroleum ether, v/v) as described above except that the
organic phase was successively washed with brine and water.
Nucleoside 2V (0.42 g, 83%) was obtained as a yellow solid material:
Rf = 0.3 (80% EtOAc in petroleum ether, v/v); MALDI-HRMS m/z
745.2225 ([M + Na]+, C35H34F3N6O8·Na+, calcd 745.2204); 1H NMR
(DMSO-d6) δ 11.40 (d, 1H, ex, J = 2.0 Hz, H3), 10.02 (t, 1H, J = 6.0
Hz, NHCOCF3), 8.01 (s, 1H, Tz), 7.81 (d, 1H, J = 8.0 Hz, H6), 7.43−
7.22 (m, 9H, DMTr), 6.93−6.88 (m, 4H, DMTr), 6.42 (d, 1H, J = 4.5
Hz, H1′), 5.79 (d, 1H, ex, J = 6.0 Hz, 3′−OH), 5.50 (dd, 1H, J = 7.0
Hz, 4.5 Hz, H2′), 5.45 (dd, 1H, J = 8.0 Hz, 2.0 Hz, H5), 4.52 (m, 1H,
H3′), 4.47 (d, 2H, J = 5.5 Hz, CH2NHCO), 4.24−4.20 (m, 1H, H4′),
3.75 (s, 6H, CH3O), 3.38−3.30 (m, 2H, H5′ − partial overlap with
6.55 (d, 1H, J = 5.0 Hz, H1′), 5.90 (d, 1H, ex, J = 5.0 Hz, 3′−OH),
5.68 (dd, 1H, J = 7.0 Hz, 5.0 Hz, H2′), 5.53 (dd, 1H, J = 8.0 Hz, 2.0
Hz, H5), 4.64−4.58 (m, 1H, H3′), 4.31−4.26 (m, 1H, H4′), 3.74 (s,
6H, CH3O), 3.40−3.30 (m, 2H, H5′); 13C NMR (DMSO-d6) δ 188.3,
162.9, 158.1, 150.2, 147.2, 144.6, 140.8 (C6), 135.4, 135.2, 133.0,
131.8, 131.5 (Tz), 130.6, 130.0, 129.74 (DMTr), 129.72 (DMTr),
129.4 (Py), 129.1 (Py), 128.9, 127.9, 127.84 (DMTr), 127.80 (Py),
127.7 (DMTr), 127.2 (Py), 126.8 (Py), 126.7 (DMTr), 126.5 (Py),
126.1 (Py), 124.01 (Py), 123.98 (Py), 123.8, 123.5, 113.2 (DMTr),
102.0 (C5), 87.4 (C1′), 85.8, 83.3 (C4′), 69.0 (C3′), 65.0 (C2′), 63.0
(C5′), 55.0 (CH3O).
5′-O-(4,4′-Dimethoxytrityl)-2′-C-[4-{2-(pyrene-1-yl)ethyl}-
1H-1,2,3-triazol-1-yl]-2′-deoxyuridine (2Y). Nucleoside 1 (0.34 g,
0.60 mmol), 4-(pyren-1-yl)-but-1-yne Ay (160 mg, 0.63 mmol),
sodium ascorbate (0.25 g, 1.25 mmol), CuSO4·5H2O (31 mg, 0.12
mmol), and THF/t-BuOH/H2O (10 mL) were mixed, reacted (2 h at rt),
worked up, and purified (50−100% EtOAc in petroleum ether,
v/v) as described above to provide nucleoside 2Y (0.33 g, 67%) as a
white solid material: Rf = 0.3 (80% EtOAc in petroleum ether, v/v);
MALDI-HRMS m/z 848.3046 ([M + Na]+, C50H43N5O7·Na+, calcd
1
848.3055); H NMR (DMSO-d6) δ 11.44 (s, 1H, ex, NH), 8.40 (d,
1H, J = 9.0 Hz, Py), 8.30−8.19 (m, 4H, Py), 8.13 (ap s, 2H, Py), 8.06
(t, 1H, J = 8.0 Hz, Py), 8.01 (s, 1H, Tz), 7.95 (d, 1H, J = 8.0 Hz, Py),
7.82 (d, 1H, J = 8.0 Hz, H6), 7.44−7.41 (m, 2H, DMTr), 7.36−7.23
(m, 7H, DMTr), 6.94−6.90 (m, 4H, DMTr), 6.44 (d, 1H, J = 5.0 Hz,
H1′), 5.79 (d, 1H, ex, J = 6.0 Hz, 3′−OH), 5.49−5.45 (m, 2H, H5,
H2′), 4.54−4.49 (m, 1H, H3′), 4.27−4.22 (m, 1H, H4′), 3.75 (s, 6H,
CH3O), 3.72−3.66 (m, 2H, CH2CH2), 3.40−3.30 (m, 2H, H5′),
3.19−3.14 (m, 2H, CH2CH2); 13C NMR (DMSO-d6) δ 162.8, 158.12,
158.11, 150.2, 145.8, 144.6, 140.5 (C6), 135.6, 135.4, 135.1, 130.8,
130.3, 129.7 (DMTr), 129.4, 128.0, 127.8 (DMTr), 127.7 (DMTr),
127.5 (Py), 127.4 (Py), 127.3 (Py), 126.7 (DMTr), 126.5 (Py), 126.1
(Py), 124.93 (Py), 124.88 (Py), 124.8 (Py), 124.2, 124.1, 123.4 (Tz),
123.2 (Py), 113.2 (DMTr), 102.0 (C5), 87.1 (C1′), 85.9, 83.3 (C4′),
68.9 (C3′), 64.3 (C2′), 62.9 (C5′), 55.0 (CH3O), 32.6 (CH2CH2),
27.3 (CH2CH2).
5′-O-(4,4′-Dimethoxytrityl)-2′-C-[4-(pyrene-1-yl)-
carboxamidomethyl-1H-1,2,3-triazol-1-yl]-2′-deoxyuridine
(2Z). Nucleoside 1 (0.40 g, 0.70 mmol), N-(prop-2-ynyl)pyrene-1-
carboxamide Az48 (200 mg, 0.71 mmol), sodium ascorbate (50 mg,
0.25 mmol), CuSO4·5H2O (5 mg, 0.02 mmol), and THF/t-BuOH/
H2O (5 mL) were mixed, reacted (8 h at rt), worked up, and purified
(50−100% EtOAc in petroleum ether, v/v) as described above except
that the organic phase was successively washed with brine and water.
Nucleoside 2Z (0.49 g, 83%) was obtained a yellow solid material: Rf =
0.2 (EtOAc); MALDI-HRMS m/z 877.2979 ([M + Na]+,
C50H42N6O8·Na+, calcd 877.2956); 1H NMR (DMSO-d6) δ 11.43
(s, 1H, ex, H3), 9.26 (t, 1H, ex, J = 6.0 Hz, NHCO), 8.53−8.52 (d,
1H, J = 9.5 Hz, Ar), 8.36−8.34 (m, 3H, Ar), 8.27−8.22 (m, 3H, Ar),
8.17−8.11 (m, 3H, Ar, Tz), 7.85 (d, 1H, J = 8.5 Hz, H6), 7.44−7.43
(m, 2H, DMTr), 7.35−7.24 (m, 7H, DMTr), 6.93−6.89 (m, 4H,
DMTr), 6.50 (d, 1H, J = 4.7 Hz, H1′), 5.87 (d, 1H, ex, J = 5.5 Hz, 3′−
OH), 5.56 (dd, 1H, J = 7.0 Hz, 4.7 Hz, H2′), 5.47 (d, 1H, J = 8.0 Hz,
H5), 4.71 (d, 2H, J = 6.0 Hz, CH2NHCO), 4.58−4.54 (m, 1H, H3′),
4.30−4.25 (m, 1H, H4′), 3.75 (s, 6H, CH3O), 3.41−3.32 (m, 2H,
H5′); 13C NMR (DMSO-d6) δ 168.8, 162.9, 158.13, 158.12, 150.2,
144.7, 144.6, 140.5 (C6), 135.4, 135.2, 131.6, 131.5, 130.7, 130.2,
129.8 (DMTr), 128.3 (Ar), 128.1 (Ar), 127.9 (DMTr), 127.8, 127.7
(DMTr), 127.1 (Ar), 126.8 (DMTr), 126.5 (Ar), 125.7 (Ar), 125.5
(Ar), 125.2 (Ar), 124.7 (Ar), 124.3 (Ar), 124.2 (Tz), 123.7, 123.6,
113.2 (DMTr), 102.0 (C5), 87.1 (C1′), 85.9, 83.3 (C4′), 69.0 (C3′),
64.5 (C2′), 62.9 (C5′), 55.0 (CH3O), 35.0 (CH2NHCO).
H2O); 13C NMR (DMSO-d6) δ 162.8, 158.09, 158.08, 156.2 (q, 1,3JCF
=
36 Hz, COCF3), 150.1, 144.6, 142.4, 140.5 (C6), 135.3, 135.1, 129.7
(DMTr), 127.8 (DMTr), 127.7 (DMTr), 126.7 (DMTr), 124.5 (Tz),
115.8 (q, JCF = 288 Hz, CF3), 113.2 (DMTr), 101.9 (C5), 87.1 (C1′),
85.8, 83.2 (C4′), 68.8 (C3′), 64.5 (C2′), 62.8 (C5′), 55.0 (CH3O), 34.5
(CH2NHCO); 19F-NMR (DMSO-d6) δ −74.2.
5′-O-(4,4′-Dimethoxytrityl)-2′-C-[4-(pyrene-1-yl)-1H-1,2,3-
triazol-1-yl]-2′-deoxyuridine (2W). Nucleoside 1 (0.28 g, 0.49
mmol), 1-ethynylpyrene Aw47 (130 mg, 0.58 mmol), sodium ascorbate
(200 mg, 1.00 mmol), CuSO4·5H2O (25 mg, 0.10 mmol), and THF/t-
BuOH/H2O (10 mL) were mixed, reacted (7 h at 75 °C), worked up,
and purified (40−70% EtOAc in petroleum ether, v/v) as described
above to provide nucleoside 2W (140 mg, 35%) as an off-white solid
material: Rf = 0.5 (80% EtOAc in petroleum ether, v/v); MALDI-
HRMS m/z 820.277 ([M + Na]+, C48H39N5O7·Na+, calcd 820.274);
1H NMR (DMSO-d6) δ 11.46 (d, 1H, ex, J = 1.5 Hz, NH), 8.87 (d,
1H, J = 9.0 Hz, Py), 8.80 (s, 1H, Tz), 8.41−8.33 (m, 4H, Py), 8.27 (d,
1H, J = 9.2 Hz, Py), 8.26−8.22 (m, 2H, Py); 8.12 (t, 1H, J = 7.5 Hz,
Py), 7.91 (d, 1H, J = 8.0 Hz, H6), 7.48−7.20 (m, 9H, DMTr), 6.96−
6.90 (m, 4H, DMTr), 6.65 (d, 1H, J = 5.0 Hz, H1′), 5.95 (d, 1H, ex,
J = 6.0 Hz, 3′−OH), 5.69 (dd, 1H, J = 7.0 Hz, 5.0 Hz, H2′), 5.54 (dd,
1H, J = 8.0 Hz, 1.5 Hz, H5), 4.69−4.64 (m, 1H, H3′), 4.40−4.36 (m,
1H, H4′), 3.76 (s, 6H, CH3O), 3.46−3.36 (m, 2H, H5′); 13C NMR
(DMSO-d6) δ 162.9, 158.2, 150.3, 145.7, 144.7, 140.8 (C6), 135.4,
135.2, 130.9, 130.6, 130.3, 129.78 (DMTr), 129.76 (DMTr), 128.0
(Py), 127.9 (DMTr), 127.73 (DMTr), 127.67 (Py), 127.5, 127.3 (Py),
127.0 (Py), 126.8 (DMTr), 126.4 (Py), 125.7 (Tz), 125.5 (Py),
125.16, 125.15 (Py), 125.09 (Py), 124.8 (Py), 124.3, 123.9, 113.3
(DMTr), 102.1 (C5), 87.4 (C1′), 85.9, 83.4 (C4′), 69.1 (C3′), 64.9
(C2′), 63.1 (C5′), 55.0 (CH3O).
5′-O-(4,4′-Dimethoxytrityl)-2′-C-[4-(pyrene-1-ylcarbonyl)-
1H-1,2,3-triazol-1-yl]-2′-deoxyuridine (2X). Nucleoside 1 (0.28 g,
0.49 mmol), 1-(pyren-1-yl)-prop-2-yn-1-one Ax (140 mg, 0.55 mmol),
sodium ascorbate (200 mg, 1.00 mmol), CuSO4·5H2O (25 mg, 0.10
mmol), and THF/t-BuOH/H2O (10 mL) were mixed, reacted (5 h
at rt), worked up, and purified (40−90% EtOAc in petroleum ether, v/v)
as described above to provide nucleoside 2X (0.25 g, 60%) as a yellow
solid material: Rf = 0.4 (80% EtOAc in petroleum ether, v/v); MALDI-
HRMS m/z 848.267 ([M + Na]+, C49H39N5O8·Na+, calcd 848.270);
1H NMR (DMSO-d6) δ 11.46 (br s, 1H, ex, NH), 8.96 (s, 1H, Tz),
General Phosphitylation Protocol for Preparation of 3V−3Z
(Description for ∼3 mmol Scale). The appropriate nucleoside 2
was coevaporated with anhydrous CH2Cl2 (5 mL) and redissolved in
anhydrous CH2Cl2 (reagent quantities and solvent volumes are
specified below). To this were added N,N-diisopropylethylamine
(DIPEA), 0.45 M tetrazole in CH3CN, and 2-cyanoethyl-N,N,N′,N′-
tetraisopropylphosphordiamidite (PN2 reagent). The reaction mixture
was stirred at rt until analytical TLC indicated complete conversion
8.51−8.28 (m, 8H, Py), 8.17 (t, 1H, J = 7.5 Hz, Py), 7.83 (d, 1H, J =
8.0 Hz, H6), 7.44−7.21 (m, 9H, DMTr), 6.93−6.88 (m, 4H, DMTr),
13
dx.doi.org/10.1021/jo201845z|J. Org. Chem. 2012, 77, 5−16

The Journal of Organic Chemistry
Featured Article
(reaction time specified below) whereupon cold absolute EtOH (0.5
mL) was added. The reaction mixture was evaporated to dryness, and
the resulting residue was purified by silica gel column chromatography
(eluent specified below). The crude material was triturated from cold
petroleum ether to afford phosphoramidite 3 (yields specified below).
3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-
O-(4,4′-dimethoxytrityl)-2′-C-[4-(2,2,2-trifluoroacetamido-
methyl)-1H-1,2,3-triazol-1-yl]-2′-deoxyuridine (3V). Nucleoside
2V (0.29 g, 0.40 mmol), DIPEA (0.10 mL, 0.57 mmol), tetrazole in
CH3CN (0.45 M, 1.0 mL, 0.45 mmol), PN2 reagent (0.15 mL, 0.46
mmol), and anhydrous CH2Cl2 (1 mL) were mixed, reacted (3 h),
worked up, and purified (50−90% EtOAc in petroleum ether, v/v) as
described above except that (a) the reaction mixture was extracted
with EtOAc (5 mL) after addition of EtOH, followed by drying of the
organic phase over anhydrous Na2SO4 and evaporation to dryness
under reduced pressure, and (b) trituration was not performed.
Phosphoramidite 3V (0.24 g, 67%) was obtained as a white solid
material: Rf = 0.3 (5% MeOH in CH2Cl2, v/v); MALDI-HRMS m/z
945.3322 ([M + Na]+, C44H50F3N8O9·Na+, calcd 945.3283); 31P NMR
(CDCl3) δ 152.0, 149.8; 19F NMR (CDCl3) δ −75.7.
3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-
O-(4,4′-dimethoxytrityl)-2′-C-[4-(pyrene-1-yl)-1H-1,2,3-triazol-
1-yl]-2′-deoxyuridine (3W). Nucleoside 2W (230 mg, 0.29 mmol),
DIPEA (0.10 mL, 0.57 mmol), tetrazole in CH3CN (0.45 M, 1.0 mL,
0.45 mmol), PN2 reagent (0.20 mL, 0.62 mmol), and anhydrous
CH2Cl2 (2 mL) were mixed, reacted (4 h), worked up, and purified
(0−4% MeOH/CH2Cl2, v/v) as described above to afford 3W (180
mg, 62%) as a white powder: Rf = 0.35 (5% MeOH in CH2Cl2, v/v);
MALDI-HRMS m/z 1020.3855 ([M + Na]+, C57H56N7O8P·Na+, calcd
1020.3826); 31P NMR (CDCl3) δ 152.0, 150.5.
3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-
O-(4,4′-dimethoxytrityl)-2′-C-[4-(pyrene-1-ylcarbonyl)-1H-
1,2,3-triazol-1-yl]-2′-deoxyuridine (3X). Nucleoside 2X (150 mg,
0.18 mmol), DIPEA (0.10 mL, 0.57 mmol), tetrazole in CH3CN (0.45
M, 0.6 mL, 0.27 mmol), PN2 reagent (0.12 mL, 0.37 mmol), and
anhydrous CH2Cl2 (2 mL) were mixed, reacted (3.5 h), worked up,
and purified (0−4% MeOH/CH2Cl2, v/v) as described above to afford
3X (110 mg, 59%) as a yellow solid material: Rf = 0.4 (5% MeOH in
CH2Cl2, v/v); MALDI-HRMS m/z 1048.3779 ([M + Na]+,
C58H56N7O9P·Na+, calcd 1048.3775); 31P NMR (CDCl3) δ 152.4,
150.9.
3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-
O-(4,4′-dimethoxytrityl)-2′-C-[4-{2-(pyrene-1-yl)ethyl}-1H-
1,2,3-triazol-1-yl]-2′-deoxyuridine (3Y). Nucleoside 2Y (0.33 g,
0.40 mmol), DIPEA (0.10 mL, 0.57 mmol), tetrazole in CH3CN (0.45
M, 1.5 mL), PN2 reagent (0.25 mL, 0.78 mmol), and anhydrous
CH2Cl2 (2 mL) were mixed, reacted (3.5 h), worked up, and purified
(0−4% MeOH/CH2Cl2, v/v) as described above to afford 3Y (210
mg, 51%) as a white powder: Rf = 0.45 (5% MeOH in CH2Cl2, v/v);
MALDI-HRMS m/z 1048.4147 ([M + Na]+, C59H60N7O8P·Na+, calcd
1048.4139); 31P NMR (CDCl3) δ 151.6, 150.6.
used. A ∼50-fold molar excess of modified phosphoramidites in
anhydrous acetonitrile (at 0.05 M) was used during hand-couplings
using the conditions specified in the article. Moreover, extended
oxidation (45 s) was employed during hand-couplings. Cleavage from
solid support and removal of protecting groups was accomplished upon
treatment with 32% aqueous ammonia (55 °C, 20 h). Purification of all
modified ONs was performed by ion-pair reverse-phase HPLC as
described below followed by detritylation (80% aqueous AcOH) and
precipitation from acetone (−18 °C for 12−16 h).
Purification of crude ONs was performed on a HPLC system
equipped with an XTerra MS C18 precolumn (10 μm, 7.8 × 10 mm)
and an XTerra MS C18 column (10 μm, 7.8 × 150 mm) using a 0.05
mM TEAA (triethylammonium acetate) buffer: 25% water/acetonitrile
(v/v) gradient. The identity of synthesized ONs was established
through analysis on a quadrupole time-of-flight tandem mass
spectrometer equipped with a MALDI source (positive ion mode)
using anthranilic acid as a matrix (Table S1 in Supporting
Information), while purity (>80%) was verified by ion-pair reverse-
phase HPLC running in analytical mode.
Thermal Denaturation Studies. Concentrations of ONs were
estimated using the following extinction coefficients (OD/μmol): dG
(12.01), dA (15.20), dT (8.40), dC (7.05); rG (13.70), rA (15.40), rU
(10.00), rC (9.00); V (19.96), W (31.08), X (35.60), Y (27.62) and Z
(30.95) [values for monomers V−Z were estimated through A260
measurements of the corresponding phosphoramidites in 1% aqueous
DMSO solutions]. Each strand was thoroughly mixed and denatured
by heating to 80−85 °C followed by cooling to the starting
temperature of the experiment. Quartz optical cells with a path length
of 10 mm were used. Thermal denaturation temperatures (Tm values
[°C]) of duplexes (1.0 μM final concentration of each strand) were
measured on a UV/vis spectrophotometer equipped with a 12-cell
Peltier temperature controller and determined as the maximum of the
first derivative of the thermal denaturation curve (A260 vs T) recorded
in medium salt buffer (Tm buffer: 100 mM NaCl, 0.1 mM EDTA, and
pH 7.0 adjusted with 10 mM Na2HPO4 and 5 mM Na2HPO4). The
temperature of the denaturation experiments ranged from at least 20
°C below Tm to 20 °C above Tm. A temperature ramp of 0.5 °C/min
was used in all experiments. Reported Tm values are averages of two
experiments within ±1.0 °C.
Steady-State Fluorescence Emission Spectra. Spectra of ONs
modified with pyrene-functionalized monomers W/X/Y/Z and the
corresponding duplexes with complementary or mismatched DNA/
RNA targets were recorded in nondeoxygenated thermal denaturation
buffer (each strand 1.0 μM) using an excitation wavelength of λex
=
350 nm for W/Y/Z or λex = 400 nm for X, excitation slit 5.0 nm,
emission slit 5.0 nm, and a scan speed of 600 nm/min. Experiments
were performed at ambient temperature (∼20 °C).
Determination of Quantum Yields. Relative fluorescence
emission quantum yields (ΦF) of modified nucleic acids (SSP or
duplex) were determined using the following equation:61
3′-O-(N,N-Diisopropylamino-2-cyanoethoxyphosphinyl)-5′-
O - ( 4 , 4 ′ - d i m e t h o x y t r i t y l ) - 2 ′ - C - [ 4 - ( p y r e n e - 1 - y l ) -
carboxamidomethyl-1H-1,2,3-triazol-1-yl]-2′-deoxyuridine
(3Z). Nucleoside 2Z (0.36 g, 0.42 mmol), DIPEA (0.10 mL, 0.57
mmol), tetrazole in CH3CN (0.45 M, 1.0 mL, 0.45 mmol), PN2
reagent (0.15 mL, 0.46 mmol), and anhydrous CH2Cl2 (1 mL) were
mixed, reacted (3 h), worked up, and purified (50−90% EtOAc in
petroleum ether, v/v) as described above except that (a) the reaction
mixture was extracted with EtOAc (5 mL) after addition of EtOH,
followed by drying of the organic phase over anhydrous Na2SO4 and
evaporation to dryness under reduced pressure, and (b) trituration was
not performed. Phosphoramidite 3Z (0.28 g, 67%) was obtained as a
white solid material: Rf = 0.3 (5% MeOH in CH2Cl2, v/v); MALDI-
HRMS m/z 1077.3984 ([M + Na]+, C59H59N8O9P·Na+, calcd
1077.4035); 31P NMR (CDCl3) δ 151.9, 150.1.
where ΦF(std) is the fluorescence emission quantum yield of the
standard; α(std) is the slope of the integrated fluorescence intensity
versus optical intensity plot made for the standard; IFI(NA) is the
integrated fluorescence intensity (λ em = 360−510 nm for monomer
W/Y/Z; λem = 425−625 nm for monomer X; λem = 360−600 nm for
standards); Aex (NA) is the optical density of the sample at the utilized
excitation wavelength (λex = 350 nm for monomer W/Y/Z; λex = 400
nm for monomer X; λex = 350 nm for standards; optical densities of all
solutions at the excitation wavelengths were between 0.01 and 0.10);
n(NA) and n(std) are refractive indexes of solvents used for sample
and standard, respectively (nwater = 1.33, nethanol = 1.36, and ncyclohexane
=
1.43).
Synthesis and Purification of ONs. Synthesis of modified
oligodeoxyribonucleotides (ONs) was performed on a DNA synthesizer
using 0.2 μmol scale succinyl-linked LCAA-CPG (long chain alkyl
amine-controlled pore glass) columns with a pore size of 500 Å.
Standard protocols for incorporation of DNA phosphoramidites were
The validity of this method under our experimental setup was
ascertained by determining the quantum yield of anthracene in ethanol
with respect to 9,10-diphenylanthracene in cyclohexane (Φ F = 0.86).61
The measured value of ΦF = 0.28 is in excellent agreement with the
14
dx.doi.org/10.1021/jo201845z|J. Org. Chem. 2012, 77, 5−16

The Journal of Organic Chemistry
Featured Article
reported value of (ΦF = 0.27).62 Subsequently, the literature value for
anthracene in ethanol was used as the standard for determination of
quantum yields of SSPs and duplexes.
Selvam, C.; Kennedy, S. D.; Rozners, E. Bioorg. Med. Chem. Lett. 2011,
21, 3420−3422. (c) El-Sagheer, A. H.; Brown, T. Proc. Natl. Acad. Sci.
U.S.A. 2010, 107, 15329−15334.
(11) Chittepu, P.; Sirivolu, V. R.; Seela, F. Bioorg. Med. Chem. 2008,
16, 8427−8439.
ASSOCIATED CONTENT
* Supporting Information
■
(12) Nakahara, M.; Kuboyama, T.; Izawa, A.; Hari, Y.; Imanishi, T.;
Obika, S. Bioorg. Med. Chem. Lett. 2009, 19, 3316−3319.
(13) Østergaard, M. E.; Guenther, D. C.; Kumar, P.; Baral, B.;
Deobald, L.; Paszczynski, A. J.; Sharma, P. K.; Hrdlicka, P. J. Chem.
Commun. 2010, 46, 4929−4931.
(14) Kumar, T. S.; Wengel, J.; Hrdlicka, P. J. ChemBioChem 2007, 8,
1122−1125.
(15) Østergaard, M. E.; Kumar, P.; Baral, B.; Guenther, D. C.;
Anderson, B. A; Ytreberg, F. M.; Deobald, L.; Paszczynski, A. J.;
Sharma, P. K.; Hrdlicka, P. J. Chem.Eur. J. 2011, 17, 3157−3165.
(16) Østergaard, M. E.; Hrdlicka, P. J. Chem. Soc. Rev. 2011, DOI:
10.1039/C1CS15014F.
(17) Østergaard, M. E.; Cheguru, P.; Papasani, M. R.; Hill, R. A.;
Hrdlicka, P. J. J. Am. Chem. Soc. 2010, 132, 14221−14228.
(18) Kumar, T. S.; Madsen, A. S.; Østergaard, M. E.; Wengel, J.;
Hrdlicka, P. J. J. Org. Chem. 2008, 73, 7060−7066.
(19) Kumar, T. S.; Madsen, A. S.; Østergaard, M. E.; Sau, S. P.;
Wengel, J.; Hrdlicka, P. J. J. Org. Chem. 2009, 74, 1070−1081.
(20) Sau, S. P.; Kumar, T. S.; Hrdlicka, P. J. Org. Biomol. Chem. 2010,
8, 2028−2036.
(21) Karmakar, S.; Anderson, B. A.; Rathje, R. L.; Andersen, S.;
Jensen, T.; Nielsen, P.; Hrdlicka, P. J. J. Org. Chem. 2011, 76, 7119−
7131.
(22) Yamana, K.; Iwase, R.; Furutani, S.; Tsuchida, H.; Zako, H.;
Yamaoka, T.; Murakami, A. Nucleic Acids Res. 1999, 27, 2387−2392.
(23) Nakamura, M.; Fukunaga, Y.; Sasa, K.; Ohtoshi, Y.; Kanaori, K.;
Hayashi, H.; Nakano, H.; Yamana, K. Nucleic Acids Res. 2005, 33,
5887−5895.
(24) Kalra, N.; Babu, B. R.; Parmar, V. S.; Wengel, J. Org. Biomol.
Chem. 2004, 2, 2885−2887.
(25) Jawalekar, A. M.; Meeuwenoord, N.; Cremers, J. S. G. O.;
Overkleeft, H. S.; Marel, G. A. V. D.; Rutjes, F. P. J. T.; Delft, F. L. V.
J. Org. Chem. 2008, 73, 287−290.
(26) Jørgensen, A. S.; Shaikh, K. I.; Enderlin, G.; Ivarsen, E.; Kumar,
S.; Nielsen, P. Org. Biomol. Chem. 2011, 9, 1381−1388.
(27) Kiviniemi, A.; Virta, P.; Drenichev, M. S.; Mikhailov, S. N.;
S
NMR spectra of all new compounds; MS data of modified
ONs; representative thermal denaturation profiles; Tm values
for 9-mer ONs modified with monomers V and Z; additional
thermal denaturation, fluorescence and absorbance data for
duplexes with matched/mismatched RNA targets. This material
is available free of charge via the Internet at http://pubs.acs.org.
AUTHOR INFORMATION
Corresponding Author
*E-mail: hrdlicka@uidaho.edu.
■
ACKNOWLEDGMENTS
■
We appreciate support from Award Number R01 GM088697
from the National Institute of General Medical Sciences,
National Institutes of Health. We thank Dr. Lee Deobald, Prof.
Andrzej J. Paszczynski (both EBI Murdock Mass Spectrometry
Center, Univ. Idaho), Prof. Matthew J. Morra, and Dr. Vladimir
Borek (both Division of Soil & Land Resources, Univ. Idaho) for
mass spectrometric analyses. The assistance of Dr. Andreas S.
Madsen (Technical University of Denmark) and Dr. Jakob
Magolan (University of Idaho) during preparation of the cover
illustration is greatly appreciated.
REFERENCES
■
(1) Rostovtsev, V. V.; Green, L. G.; Fokin, V. V.; Sharpless, K. B.
Angew. Chem., Int. Ed. 2002, 41, 2596−2599.
(2) Tornøe, C. W.; Christensen, C.; Meldal, M. J. Org. Chem. 2002,
67, 3057−3064.
(3) Amblard, F.; Cho, J. H.; Schinazi, R. F. Chem. Rev. 2009, 109,
4207−4220.
(4) Ustinov, A. V.; Stepanova, I. A.; Dubnyakova, V. V.; Zatsepin, T.
S.; Nozhevnikova, E. V.; Korshun, V. A. Russ. J. Bioorg. Chem. 2010, 36,
437−481.
Lonnberg, H. Bioconjugate Chem. 2011, 22, 1249−1255.
̈
(5) El-Sagheer, A. H.; Brown, T. Chem. Soc. Rev. 2010, 39, 1388−
1405.
(28) Patil, R. V.; Dekker, E. E. Nucleic Acids Res. 1990, 18, 3080−
3080.
(6) (a) Seo, T. S.; Li, Z.; Ruparel, H.; Ju, J. J. Org. Chem. 2003, 68,
609−612. (b) Gierlich, J.; Burley, G. A.; Gramlich, P. M. E.;
Hammond, D. M.; Carell, T. Org. Lett. 2006, 8, 3339−3342. (c) Seela,
F.; Sirivolu, V. R.; Chittepu, P. Bioconjugate Chem. 2008, 19, 211−224.
(d) Seela, F.; Ming, X. Helv. Chim. Acta 2008, 91, 1181−1200.
(e) Berndl, S.; Herzig, N.; Kele, P.; Lachmann, D.; Li, X.; Wolfbeis, O.
S.; Wagenknecht, H.-A. Bioconjugate Chem. 2009, 20, 558−564.
(f) Seela, F.; Ingale, S. A. J. Org. Chem. 2010, 75, 284−295. (g) Ding,
P.; Wunnicke, D.; Steinhoff, H.-J.; Seela, F. Chem.Eur. J. 2010, 16,
14385−14396. (h) Jakobsen, U.; Shelke, S. A; Vogel, S.; Sigurdsson, S.
T. J. Am. Chem. Soc. 2010, 132, 10424−10428. (i) Onizuka, K.;
Shibata, A.; Taniguchi, Y.; Sasaki, S. Chem. Commun. 2011, 47, 5004−
5006. (j) Kuboyama, T.; Nakahara, M.; Yoshino, M.; Cui, Y.; Sako, T.;
Wada, Y.; Imanishi, T.; Obika, S.; Watanabe, Y.; Suzuki, M.; Doi, H.
Bioorg. Med. Chem. 2011, 19, 249−255. (k) Mercier, F.; Paris, J.;
Kaisin, G.; Thonon, D.; Flagothier, J.; Teller, N.; Lemaire, C.; Luxen,
A. Bioconjugate Chem. 2011, 22, 108−114.
(7) Burley, G. A.; Gierlich, J.; Mofid, M. R.; Nir, H.; Tal, S.; Eichen,
Y.; Carell, T. J. Am. Chem. Soc. 2006, 128, 1398−1399.
(8) Kumar, R.; El-Sagheer, A.; Tumpane, J.; Lincoln, P.;
Wilhelmsson, L. M.; Brown, T. J. Am. Chem. Soc. 2007, 129, 6859−
6864.
(9) Xiong, H.; Seela, F. J. Org. Chem. 2011, 76, 5584−5597.
(10) (a) Isobe, H.; Fujino, T.; Yamazaki, N.; Guillot-Nieckowski, M.;
Nakamura, E. Org. Lett. 2008, 10, 3729−3732. (b) Mutisya, D.;
(29) Nichols, R.; Andrews, P. C.; Zhang, P.; Bergstrom, D. E. Nature
1994, 369, 492−493.
(30) Loakes, D.; Brown, D. M.; Linde, S.; Hill, F. Nucleic Acids Res.
1995, 23, 2361−2366.
(31) Chizhikov, V.; Wagner, M.; Ivshina, A.; Hoshino, Y.; Kapikian,
A. Z.; Chumakov, K. J. Clin. Microbiol. 2002, 40, 2398−2407.
(32) Too, K.; Loakes, D. Universal Base Analogues and their
Application to Biotechnology. In Modified Nucleosides: In Biochemistry,
Biotechnology and Medicine, Herdewijn, P., Ed.; Wiley-VCH:
Weinheim, Germany, 2008; pp 277−303.
(33) Loakes, D.; Brown, D. M. Nucleic Acids Res. 1994, 22, 4039−
4043.
(34) Berger, M.; Wu, Y.; Ogawa, A. K.; McMinn, D. L.; Schultz, P.
G.; Romesberg, F. E. Nucleic Acids Res. 2000, 28, 2911−2914.
(35) Matray, T. J.; Kool, E. T. J. Am. Chem. Soc. 1998, 120, 6191−
6192.
(36) Babu, B. R.; Wengel, J. Chem. Commun. 2001, 2114−2115.
(37) Raunak.; Babu, B. R.; Parmar, V. S.; Harrit, N. H.; Wengel, J.
Org. Biomol. Chem. 2004, 2, 80−89.
(38) MacKinnon, K. F.; Qualley, D. F.; Woski, S. A. Tetrahedron Lett.
2007, 48, 8074−8077.
(39) Seela, F.; Chen, Y. Nucleic Acids Res. 1995, 23, 2499−2505.
(40) Acedo, M.; De Clercq, E.; Eritja, R. J. Org. Chem. 1995, 60,
6262−6269.
15
dx.doi.org/10.1021/jo201845z|J. Org. Chem. 2012, 77, 5−16

The Journal of Organic Chemistry
Featured Article
́
(41) Cubero, E.; Guimil-Garcia, R.; Luque, F. J.; Eritja, R.; Orozco, M.
̈
Nucleic Acids Res. 2001, 29, 2522−2534.
(42) Seela, F.; Debelak, H. Nucleic Acids Res. 2000, 28, 3224−3232.
(43) Koller, A. N.; Bozilovic, J.; Engels, J. W.; Gohlke, H. Nucleic
Acids Res. 2010, 38, 3133−3146.
(44) For a successful example, see: Kohler, O.; Jarikote, D. V.; Seitz, O.
̈
ChemBioChem 2005, 6, 69−77.
(45) Patra, A.; Richert, C. J. Am. Chem. Soc. 2009, 131, 12671−
12681.
(46) Trybulski, E. J.; Zhang, J.; Kramss, R. H.; Mangano, R. M.
J. Med. Chem. 1993, 36, 3533−3541.
(47) Liu, F.; Tang, C.; Chen, Q.-Q.; Li, S.-Z.; Wu, H.-B.; Xie, L.-H.;
Peng, B.; Wei, W.; Cao, Y.; Huang, W. Org. Electron. 2009, 10, 256−
265.
(48) Okamoto, A.; Kanatani, K.; Saito, I. J. Am. Chem. Soc. 2004, 126,
4820−4827.
(49) Sanghvi, Y. S.; Hoke, G. D.; Freier, S. M.; Zounes, M. C.;
Gonzalez, C.; Cummins, L.; Sasmor, H.; Cook, P. D. Nucleic Acids Res.
1993, 21, 3197−3203.
(50) Oliver, J. S.; Parker, K. A.; Suggs, J. W. Org. Lett. 2001, 3, 1977−
1980.
(51) Millican, T. A.; Mock, G. A.; Chauncey, M. A.; Patel, T. P.;
Eaton, M. A. W.; Gunning, J.; Cutbush, S. D.; Neidle, S.; Mann, J.
Nucleic Acids Res. 1984, 12, 7435−7453.
(52) Langenegger, S. M.; Haner, R. ChemBioChem 2005, 6, 848−
̈
851.
(53) Nakano, S.; Uotani, Y.; Uenishi, K.; Fujii, M.; Sugimoto, N.
Nucleic Acids Res. 2005, 33, 7111−7119.
́
(54) Verhagen, C.; Bryld, T.; Raunkjær, M.; Vogel, S.; Buchalova, K.;
Wengel, J. Eur. J. Org. Chem. 2006, 2538−2548.
(55) Dougherty, G.; Pilbrow, J. R. Int. J. Biochem. 1984, 16, 1179−
1192.
(56) Manoharan, M.; Tivel, K. L.; Zhao, M.; Nafisi, K.; Netzel, T. L.
J. Phys. Chem. 1995, 99, 17461−1747.
(57) Seo, Y. J.; Ryu, J. H.; Kim, B. H. Org. Lett. 2005, 7, 4931−4933.
(58) Jiang, Y. L.; Kwon, K.; Stivers, J. T. J. Biol. Chem. 2001, 276,
42347−42354.
(59) Beuck, C.; Singh, I.; Bhattacharya, A.; Hecker, W.; Parmar, V. S.;
Seitz, O.; Weinhold, E. Angew. Chem., Int. Ed. 2003, 42, 3958−3960.
(60) Fleming, D. A.; Thode, C. J.; Williams, M. E. Chem. Mater.
2006, 18, 2327−2334.
(61) Morris, J. V.; Mahaney, M. A.; Huber, J. R. J. Phys. Chem. 1976,
80, 969−974.
(62) Melhuish, W. H. J. Phys. Chem. 1961, 65, 229−235.
16
dx.doi.org/10.1021/jo201845z|J. Org. Chem. 2012, 77, 5−16