C5-functionalized DNA, LNA, and α- L -LNA: Positional control of polarity-sensitive fluorophores leads to improved SNP-Typing

Article abstract of DOI:10.1002/chem.201002109

Single nucleotide polymorphisms (SNPs) are important markers in disease genetics and pharmacogenomic studies. Oligodeoxyribonucleotides (ONs) modified with 5-[3-(1-pyrenecarboxamido)propynyl]-2′-deoxyuridine monomer X enable detection of SNPs at non-strin

Full text of DOI:10.1002/chem.201002109

FULL PAPER
DOI: 10.1002/chem.201002109
C5-Functionalized DNA, LNA, and a-l-LNA: Positional Control of Polarity-
Sensitive Fluorophores Leads to Improved SNP-Typing**
Michael E. Østergaard,[a] Pawan Kumar,[a, b] Bharat Baral,[a] Dale C. Guenther,[a]
Brooke A. Anderson,[a] F. Marty Ytreberg,[c] Lee Deobald,[d] Andrzej J. Paszczynski,[d]
Pawan K. Sharma,[b] and Patrick J. Hrdlicka*[a]
Abstract: Single nucleotide polymorph-
isms (SNPs) are important markers in
disease genetics and pharmacogenomic
monomers Y or Z result in a) larger in-
creases in fluorescence intensity upon
hybridization to complementary DNA,
b) formation of more brightly fluores-
cent duplexes due to markedly larger
fluorescence emission quantum yields
(FF =0.44–0.80) and pyrene extinction
coefficients, and c) improved optical
discrimination of SNPs in DNA targets.
Optical spectroscopy studies suggest
that the nucleobase moieties of mono-
mers X–Z adopt anti and syn confor-
mations upon hybridization with
matched and mismatched targets, re-
spectively. The polarity-sensitive 1-pyr-
enecarboxamido fluorophore is, there-
by, either positioned in the polar major
groove or in the hydrophobic duplex
core close to quenching nucleobases.
Calculations suggest that the bicyclic
skeletons of LNA and a-l-LNA mono-
mers Y and Z influence the glycosidic
torsional angle profile leading to al-
tered positional control and photophys-
ical properties of the C5-fluorophore.
studies.
Oligodeoxyribonucleotides
(ONs) modified with 5-[3-(1-pyrenecar-
boxamido)propynyl]-2’-deoxyuridine
monomer X enable detection of SNPs
at non-stringent conditions due to dif-
ferential fluorescence emission of
matched versus mismatched nucleic
acid duplexes. Herein, the thermal de-
naturation and optical spectroscopic
characteristics of monomer X are com-
pared to the corresponding locked nu-
cleic acid (LNA) and a-l-LNA mono-
mers Y and Z. ONs modified with
Keywords: bicyclic nucleotides
·
BNA · DNA targeting · pyrene ·
single nucleotide polymorphisms
Introduction
bility between duplexes of probes and complementary or
SNP-containing targets.[1,2] Moreover, these multistep proto-
cols often necessitate stringent control of assay conditions
(e.g., temperature, ionic strength). As a result, there has
been a major thrust to develop alternative SNP-typing ap-
proaches, which are operationally more simple and cost-effi-
cient. Examples include modified molecular beacons,[3] dual
probes,[4] quenched autoligation probes,[5] intercalator-modi-
fied probes,[6] charge transfer based approaches,[7] and base-
discriminating fluorescent (BDF) probes.[8] Oligodeoxyribo-
nucleotides (ONs) modified with 5-[3-(1-pyrenecarboxami-
do)propynyl]-2’-deoxyuridine monomer X (Figure 1) have
emerged as particularly promising BDF probes due to their
efficient optical discrimination of complementary over mis-
matched targets and moderately high fluorescence quantum
yields,[8d] which has enabled discrimination of SNPs in
human breast cancer cell lines at 50 nm target concentra-
tion.[9] Molecular modeling and photophysical studies sug-
gest that the polarity-sensitive 3-(1-pyrenecarboxamido)pro-
pynyl moiety of monomer X (Figure 1) intercalates into the
hydrophobic base stack upon hybridization with mismatched
targets resulting in fluorescence quenching, while it points
toward the polar major groove in duplexes with complemen-
tary DNA targets resulting in high fluorescence.[8d] These
differences in binding modes are suggested to correlate with
changes in the glycosidic torsion angle (O4’-C1’-N1-C2)
from anti to syn ranges.
Single nucleotide polymorphisms (SNPs) are the most fre-
quently occurring genetic variation in the human genome
(>9 million SNPs, one SNP per 1000 base pairs).[1] SNPs
often result in phenotypic changes, and are accordingly im-
portant markers in disease genetics and pharmacogenomic
studies. The most established SNP genotyping technologies
are enzyme-based or rely on small differences in thermosta-
[a] Dr. M. E. Østergaard, Dr. P. Kumar, B. Baral, D. C. Guenther,
B. A. Anderson, Prof. P. J. Hrdlicka
Department of Chemistry, University of Idaho
P.O. Box 442343, Moscow, ID 83844-2343 (USA)
Fax : (+1)208-885-6173
E-mail: hrdlicka@uidaho.edu
[b] Dr. P. Kumar, Prof. P. K. Sharma
Department of Chemistry, Kurukshetra University
Kurukshetra 136119 (India)
[c] Prof. F. M. Ytreberg
Department of Physics, University of Idaho
P. O. Box 440903, Moscow, ID 83844-0903 (USA)
[d] Dr. L. Deobald, Prof. A. J. Paszczynski
Environmental Biotechnology Institute, University of Idaho
P. O. Box 441052, Moscow, ID 83844-1052 (USA)
[**] LNA=locked nucleic acid; SNP=single nucleotide polymorphism.
Supporting information for this article is available on the WWW
under http://dx.doi.org/10.1002/chem.201002109.
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Results and Discussion
Synthesis: The Sonogashira reaction between known 5-iodo-
LNA uridine derivative 1Y[10c] and 1-pyrenylpropargyl-
AHCTUNGTRENNUNG
amide[8d] afforded C5-functionalized alcohol 2Y in 74%
yield (Scheme 1). Subsequent O3’-phosphitylation using
standard conditions afforded phosphoramidite 3Y in 61%
yield. The corresponding C5-functionalized a-l-LNA phos-
phoramidite 3Z, that is, the first example of this class of
LNAs, was obtained in an equivalent manner from 1Z
(Scheme 2; the synthesis of this intermediate will be pre-
sented elsewhere), while the 2’-deoxyuridine analogue was
prepared as reported in the literature.[8d] The identity of all
new compounds was fully ascertained by NMR spectroscopy
(1H, 13C, 31P, COSY and HSQC) and HRMS, while purity
was determined by 1D NMR spectroscopy (see the Support-
ing Information)
The phosphoramidites were used in automated solid-
phase DNA synthesis (0.2 mmol scale) to incorporate mono-
mers X, Y and Z into the center of 13-mer ONs that have
been previously used to study BDF probes.[8d] The nucleo-
tides flanking the X/Y/Z monomers were systematically
varied to explore the influence of neighbouring nucleotides
on biophysical properties (ON5–ON16). Coupling yields of
>98% were observed during incorporation of unmodified
monomers as well as for phosphoramidites 3X/3Y/3Z using
extended coupling times (15 min) and 4,5-dicyanoimidazole
as an activator (see the Supporting Information) The identi-
ty and purity of the ONs was verified by MALDI-TOF MS
analysis (see Table S2 in the Supporting Information) and
RP-HPLC (>80%), respectively.
Thermal denaturation studies: The thermostability of du-
plexes between ON5–ON16 and complementary or mis-
matched DNA targets was studied by UV thermal denatura-
tion experiments using medium salt buffer conditions
Figure 1. Chemical structures of 5-[3-(1-pyrenecarboxamido)propynyl]-2’-
deoxyuridine, LNA uridine and a-l-LNA uridine monomers X–Z studied
herein. Pyr=pyren-1-yl. Illustration of suggested interaction between H6
and furanose hydrogens.
AHCTUNGTRENNUNG
([Na+]=110 mm) and compared to the corresponding un-
modified DNA duplexes (Table 1). The UV thermal denatu-
ration curves of all modified DNA duplexes exhibit smooth
sigmoidal monophasic transitions (see Figures S1–S3 in the
Supporting Information). Less pronounced hyperchromicity
was observed in duplexes modified with C5-functionalized
LNA monomer Y or a-l-LNA monomer Z, which indicates
less efficient p–p stacking between nucleobases in the
duplex.[14] Incorporation of the known 5-[3-(1-pyrenecarbox-
amido)propynyl]-2’-deoxyuridine monomer X[8d] into ONs
(ON5–ON8) significantly decreases the thermal denatura-
tion temperatures (Tm) of duplexes with DNA complements
(DTm between ꢀ1.58C and ꢀ6.08C, Table 1). Interestingly,
incorporation of the corresponding C5-functionalized LNA
or a-l-LNA building blocks into ONs results in similar de-
stabilization of duplexes although with reduced sequence
variability (DTm between ꢀ2.0 and ꢀ4.58C for monomer Y,
and between ꢀ2.5 and ꢀ4.08C for monomer Z, Table 1).
Thus, the well-established stabilizing effects of conventional
LNA[11] and a-l-LNA[12] monomers appear to be fully com-
promised by the 3-(1-pyrenecarboxamido)propynyl moiety
As part of our ongoing interest in functionalized var-
iants[10] of LNA (Locked Nucleic Acid)[11] and a-l-LNA,[12]
we set out to synthesize and characterize the corresponding
C5-functionalized LNA and a-l-LNA monomers Y and Z in
a comparative study with monomer X (Figure 1). We hy-
pothesized that the extreme sugar puckering of the bicyclic
LNA and a-l-LNA nucleotides[13] would a) influence the
anti to syn rotational profile due to steric hindrance between
H6 and sugar protons (Figure 1), b) result in higher posi-
tional control of the polarity-sensitive 3-(1-pyrenecarboxa-
mido)propynyl fluorophore, and c) possibly lead to im-
proved SNP-typing properties.
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SNP Discrimination with LNA
FULL PAPER
ON1–ON4 exhibit the expected
specificity patterns, that is, a)
formation of duplexes with sub-
stantially reduced thermostabil-
ity, and b) less efficient discrim-
ination of T:G mismatches
compared to T:C and T:T mis-
matches (Table 1). Comparison
with modified ON5–ON16 in
the same sequence contexts re-
veals that the 3-(1-pyrenecar-
boxamido)propynyl moiety at
the C5 position markedly de-
creases target specificity as the
following order is observed:
thymidine [highest specificity]>
a-l-LNA monomer Z> LNA
monomer Y> DNA monomer
X (e.g., compare mismatch DTm
values for ON2, ON6, ON10,
and ON14, Table 1). This lends
support for the hypothesis that
the pyrene moiety intercalates
upon hybridization with mis-
matched targets, as decreased
mismatch specificity often is ob-
served for monomers with in-
tercalating units.[10b,15] The im-
proved mismatch discrimination
of LNA monomer Y and a-l-
LNA monomer Z relative to
DNA monomer X parallels the
well-established enhanced mis-
Scheme 1. Synthesis of monomer Y. DMTr=4,4’-dimethoxytrityl.
match
discrimination
of
LNA[11a,16] and a-l-LNA,[10b,12]
although the underlying molec-
ular mechanism leading to this
effect is unknown.
Optical spectroscopy studies:
Three characteristics are of par-
ticular importance in the design
of SNP probes: a) the relative
increase in fluorescence intensity
upon hybridization to comple-
mentary nucleic acid targets,
since excess probe cannot be
washed out in homogeneous
Scheme 2. Synthesis of monomer Z.
assays, b) the brightness of the
resulting target duplexes, de-
fined as the product of the ex-
at the C5 position. This contrasts trends with C5-functional-
ized LNA carrying non-aromatic moieties, which generally
are very well tolerated in nucleic acid duplexes.[10c]
Next, the Watson–Crick specificity of ON5–ON16 was
evaluated by using DNA targets with mismatched nucleo-
tides opposite of the site of modification. Reference strands
tinction coefficient of the fluorophore at the applied excita-
tion wavelength eex and the fluorescence emission quantum
yield FF, since this influences detection limits, and c) the op-
tical discrimination of singly mismatched nucleic acid targets,
since this determines the robustness of the SNP-typing
method. To fully evaluate these characteristics and gain ad-
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P. J. Hrdlicka et al.
Table 1. Thermal denaturation temperatures (Tm values) of duplexes between ON1–ON16 and complementary
(B=A) or mismatched DNA targets.[a]
ONs modified with DNA mo-
nomer X (ON5–ON8) and com-
plementary DNA is accompa-
nied with 2.4- to 9.6-fold in-
creases in fluorescence intensity
(see values above black histo-
grams in Figure 4 upper panel)
and high fluorescence quantum
yields (FF =0.33–0.58, Table 2).
In concert, these observations
suggest decreased electronic in-
teractions between pyrene and
quenching nucleobase moieties
upon duplex formation.[19] ONs
modified with LNA monomer
Y (ON9–ON12) display similar
hybridization-induced increases
Tm (DTm)
[8C][b]
Mismatch DTm
[8C][c]
ON
Sequences
B=A[b]
C[c]
G[c]
T[c]
1
2
3
4
5’-CG CAA ATA AAC GC
5’-CG CAA CTC AAC GC
5’-CG CAA GTG AAC GC
5’-CG CAA TTT AAC GC
48.5
55.5
55.0
48.5
ꢀ10.0
ꢀ13.5
ꢀ13.0
ꢀ11.0
ꢀ5.0
ꢀ7.0
ꢀ9.5
ꢀ9.0
ꢀ9.0
ꢀ9.0
ꢀ10.0
ꢀ11.0
5
6
7
8
5’-CG CAA AXA AAC GC
5’-CG CAA CXC AAC GC
5’-CG CAA GXG AAC GC
5’-CG CAA TXT AAC GC
45.0 (ꢀ3.5)
54.0 (ꢀ1.5)
49.0 (ꢀ6.0)
44.0 (ꢀ4.5)
ꢀ4.5
ꢀ8.0
ꢀ3.5
ꢀ5.0
ꢀ2.0
ꢀ4.0
ꢀ7.0
ꢀ4.0
ꢀ3.0
ꢀ5.5
ꢀ4.5
ꢀ3.5
9
5’-CG CAA AYA AAC GC
5’-CG CAA CYC AAC GC
5’-CG CAA GYG AAC GC
5’-CG CAA TYT AAC GC
45.5 (ꢀ3.0)
53.5 (ꢀ2.0)
51.5 (ꢀ3.5)
44.0 (ꢀ4.5)
ꢀ5.5
ꢀ9.0
ꢀ3.5
ꢀ7.5
ꢀ3.5
ꢀ4.5
ꢀ11.5
ꢀ6.5
ꢀ4.5
ꢀ7.0
ꢀ6.5
ꢀ6.0
10
11
12
13
14
15
16
5’-CG CAA AZA AAC GC
5’-CG CAA CZC AAC GC
5’-CG CAA GZG AAC GC
5’-CG CAA TZT AAC GC
44.5 (ꢀ4.0)
52.5 (ꢀ3.0)
52.5 (ꢀ2.5)
45.0 (ꢀ3.5)
ꢀ7.5
ꢀ11.0
ꢀ8.0
ꢀ9.5
ꢀ6.5
ꢀ2.5
ꢀ6.0
ꢀ8.0
ꢀ3.0
in
fluorescence
intensity
ꢀ6.5
(Figure 4 middle panel) but
form duplexes with even higher
quantum yields, particularly in
the CYC- and GYG-sequence
contexts (ON10 and ON11,
Table 2). Interestingly, ONs
modified with a-l-LNA mono-
mer Z display larger hybridiza-
tion-induced increases in fluo-
ꢀ12.0
ꢀ10.0
[a] Tm values measured as maximum of first derivative plot of melting curves (A260 vs. T) recorded in medium
salt buffer solution ([Na+]=110 mm, [Clꢀ]=100 mm, pH 7.0 (NaH2PO4/Na2HPO4), EDTA=0.2 mm) using
1.0 mm concentration of each strand. Tm values are averages of at least two measurements. [b] (DTm)=change
in Tm value relative to unmodified reference duplex for example, ON5:DNA versus ON1:DNA. [c] Mismatch
DTm =difference in Tm value between mismatched duplex and complementary duplex; mismatched sequences:
3’-GC GTT TBT TTG CG-5ꢁ (for ON1/ON5/ON9/ON13), 3’-GC GTT GBG TTG CG-5ꢁ (for ON2/ON6/
ON10/ON14), 3’-GC GTT CBC TTG CG-5ꢁ (for ON3/ON7/ON11/ON15) and 3’-GC GTT ABA TTG CG-5ꢁ
(for ON4/ON8/ON12/ON16) where B is A, C, G, and T.
ditional insight into the binding mode of the fluorophore,
we recorded absorption, steady-state fluorescence emission
(lex =344 nm) and fluorescence excitation (lem =404 nm)
spectra of single stranded probes (SSPs) ON5–ON16 and of
the corresponding duplexes with complementary or mis-
matched DNA targets. Low experimental temperatures
(58C) were chosen to maximize strand hybridization. Deox-
ygenation was deliberately not applied to the samples since
the scope of the work was to determine fluorescence en-
hancement under aerated conditions prevailing in bioassays.
In addition, cross-calibrated fluorescence emission quantum
yields (FF) were determined relative to pyrenebutanoic acid
in methanol (FF =0.065)[17] and 9,10-diphenylanthracene in
cyclohexane (FF =0.95)[18] following established protocols
(see the Supporting Information).[17]
Figure 2. Absorption spectra of ON15 and duplexes with matched or mis-
matched DNA targets (mismatched nucleotide opposite of incorporation
site mentioned in parenthesis). Buffers and conditions are as described
for Tm experiments, T=58C.
Hybridization of ON5–ON16 to complementary DNA is
accompanied by a) hypsochromic shifts in absorption and
excitation maxima of the pyrene moiety of 3–7 nm from ap-
proximately 345–350 nm to approximately 342–346 nm
(Figure 2, as well as Figures S4 and S5, and Table S3 and S4
in the Supporting Information), b) increases in extinction
coefficients (hyperchromic shifts) that appear to be most
pronounced for a-l-LNA monomer Z and least pronounced
for DNA monomer X (Table 2), and c) formation of duplex-
es that exhibit broad fluorescence emission maxima at ap-
rescence intensity in the AZA- and TZT-contexts than cor-
responding ONs modified with X or Y monomers (e.g.,
compare ON8/ON12/ON16, Figure 4 lower panel), while
similar increases are observed in the other sequence con-
texts. In addition, duplexes display the highest observed
quantum yields in this study (FF =0.50–0.80, Table 2) which
suggests that the fluorophore experiences even fewer
quenching interactions with nucleobases. This is most likely
a consequence of a more rigid positioning of the fluoro-
proximately 402 nm with
a shoulder at approximately
387 nm (Figure 3 as well as Figures S6–S8 and Table S5 in
the Supporting Information). Duplex formation between
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SNP Discrimination with LNA
FULL PAPER
Table 2. Fluorescence quantum yields (FF) and extinction coefficients e344 of ON5–ON16 in the absence (SSP)
and TXT-sequence contexts,
which demonstrates the need
for nearby guanine moieties to
quench pyrene monomer fluo-
rescence[15b,22] and ensure effi-
cient SNP discrimination. ONs
modified with LNA monomer
Y generally display improved
SNP discrimination in the
CYC- and GYG-sequence con-
texts (Im/Imm =7.3–15.9 and 8.3–
41.9 for ON10 and ON11, re-
or presence of matched or mismatched DNA targets.[a]
FF (e344nm [cmꢀ1 mmꢀ1])
ON
Sequences
SSP
B=A
C
G
T
5
6
7
8
5’-CG CAA AXA AAC GC
5’-CG CAA CXC AAC GC
5’-CG CAA GXG AAC GC
5’-CG CAA TXT AAC GC
0.21 (22)
0.16 (19)
0.04 (19)
0.17 (21)
0.44 (24)
0.43 (24)
0.33 (20)
0.58 (23)
0.31 (17)
0.08 (17)
0.04 (17)
0.32 (18)
0.10 (17)
0.08 (16)
0.05 (15)
0.15 (17)
0.16 (16)
0.09 (18)
0.03 (17)
0.12 (16)
9
5’-CG CAA AYA AAC GC
5’-CG CAA CYC AAC GC
5’-CG CAA GYG AAC GC
5’-CG CAA TYT AAC GC
0.32 (19)
0.27 (20)
0.04 (25)
0.31 (20)
0.48 (23)
0.67 (29)
0.44 (29)
0.61 (28)
0.47 (17)
0.14 (18)
0.02 (20)
0.62 (17)
0.16 (15)
0.09 (16)
0.07 (20)
0.17 (18)
0.45 (16)
0.12 (17)
0.04 (20)
0.31 (16)
10
11
12
13
14
15
16
5’-CG CAA AZA AAC GC
5’-CG CAA CZC AAC GC
5’-CG CAA GZG AAC GC
5’-CG CAA TZT AAC GC
0.17 (11)
0.23 (20)
0.08 (23)
0.28 (12)
0.53 (16)
0.61 (30)
0.50 (31)
0.80 (20)
0.22 (8)
0.19 (17)
0.05 (20)
0.44 (9)
0.25 (11)
0.24 (23)
0.27 (27)
0.40 (14)
0.07 (9)
0.10 (14)
0.05 (16)
0.17 (9)
spectively,
Figure 4
middle
panel) but less efficient discrim-
ination in the AYA- and TYT-
sequence contexts. By contrast,
ONs modified with a-l-LNA
monomer Z (ON13–ON16) dis-
play distinctly different fluores-
[a] Conditions as described in footnote of Table 1. lex =344 nm, T=58C, 1.0 mm concentration of each strand.
phore in the major groove. Similar observations have been
reported for LNA analogues linked to fluorophores that
point to the minor groove.[10f,20]
cence trends than ON5–ON12, for example, the Z:G-mis-
matches are the least efficiently discriminated mismatches
(Figure 4 lower panel). This behaviour is surprising given
the aforementioned efficiency of guanine to quench pyrene
monomer fluorescence[15b,22] and that other SNP-typing
probes discriminate G-mismatches well.[8d,g,h] Moreover, im-
proved SNP discrimination is observed in the challenging
AZA- and TZT-contexts although higher discrimination fac-
tors, in particular with G-mismatches, would be desirable for
practical diagnostic applications (Im/Imm =3.4–14.1 and 2.9–
11.4 for ON13 and ON16, respectively, Figure 3 right panel
and Figure 4 lower panel).
Interestingly, the observed intensity-based mismatch dis-
crimination factors of ON5–ON16 are not fully accounted
by the differences in quantum yields between matched and
mismatched duplexes, for example, compare Im/Imm =5.8
with FF,m/FF,mm =0.61/0.19 ꢁ3.2 for ON14 versus the C-mis-
matched target (Figure 4 and Table 2, respectively). This re-
flects the fact that fluorescence brightness depends on the
emission quantum yield FF and the extinction coefficient e
of the fluorophore, which is markedly lower in mismatched
Next, the optical properties of mismatched duplexes were
studied. Pronounced bathochromic shifts of pyrene absorp-
tion (2–8 nm, Figure 2, and Table S3) and fluorescence emis-
sion maxima (up to 5 nm, Figure 3 as well as Figures S6–S8
and Table S5 in the Supporting Information) were observed
relative to matched duplexes along with large hypochromic
shifts in pyrene absorption (Table 2). This points toward in-
creased interactions between pyrene and nucleobase moiet-
ies in mismatched duplexes,[19] which is indicative of an in-
creasingly intercalative binding mode of the pyrene fluoro-
phore.[8d,10b,21]
Probes where DNA monomer X is incorporated between
flanking cytosine or guanine moieties (ON6 & ON7) display
excellent optical discrimination of SNPs (Figure 3 left panel
and Figure S7) with large discrimination factors Im/Imm of
7.4–8.4 and 9.7–12.9 (values above blue/yellow/green histo-
grams in Figure 4 upper panel, see legend for definition of
Im/Imm). Less efficient discrimination is observed in AXA-
Figure 3. Steady-state fluorescence spectra of ON7 (left), ON11 (middle) and ON15 (right) in the absence or presence of matched or mismatched DNA
targets (mismatched nucleotide opposite of incorporation site mentioned in parenthesis). Buffers and conditions are as described for Tm experiments.
lex =344 nm; T=58C, 1.0 mm concentration of each strand.
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P. J. Hrdlicka et al.
mentary or singly mismatched RNA targets. Similar trends
in duplex thermostability (see Table S6), target specificity
(see Table S6), excitation maxima (see Table S8), and hy-
bridization-induced increases in fluorescence intensity (see
Figures S11–S13) are observed as in the corresponding stud-
ies with DNA targets. However, the optical discrimination
of mismatched RNA targets is generally much less efficient
(see Figure S13) which is accompanied by higher quantum
yields for mismatched duplexes (see Table S7). These obser-
vations point toward less facile intercalation of the pyrene
moiety in mismatched RNA duplexes, possibly due to the
more compressed DNA:RNA duplex architectures[24] which
leads to less efficient nucleobase-mediated quenching.
Probes modified with monomer X or Y display similar opti-
cal discrimination of RNA-mismatches, whereas monomer Z
modified probes display slightly improved discrimination in
the TZT-context (see Figure S13, lower panel). Matched du-
plexes involving ONs modified with LNA monomer Y or a-
l-LNA monomer Z exhibit higher quantum yields than with
monomer X (see Table S7).
Integrated discussion: The results demonstrate that probes
based on LNA monomer Y offer practical advantages as
SNP-detection probes compared to the original 5-[3-(1-pyre-
necarboxamido)propynyl]-2’-deoxyuridine probes (monomer
X) in CYC- and GYG-sequence contexts (higher quantum
yields and improved optical discrimination of SNPs). Probes
modified with a-l-LNA monomer Z, on the other hand, dis-
play interesting characteristics in the challenging AZA- and
TZT-contexts (larger hybridization-induced increases, higher
quantum yields and improved optical discrimination of
SNPs). Results from optical spectroscopy studies strongly
suggest that the nucleobase moieties of monomers X-Z
adopt anti and syn conformations upon hybridization with
matched and mismatched targets, respectively. The polarity-
sensitive 1-pyrenecarboxamido fluorophore is thereby either
positioned in the polar major groove or in the hydrophobic
duplex core close to quenching nucleobases. The improved
SNP-discrimination by LNA and a-l-LNA monomers Y and
Z, is a consequence of higher emission quantum yields of
matched duplexes (monomers Y and Z), high pyrene extinc-
tion coefficients in matched duplexes (monomer Y), and
low pyrene extinction coefficients in mismatched duplexes
(monomer Z). Calculations indicate that the bicyclic skele-
tons of the LNA and a-l-LNA monomers Y and Z influence
the glycosidic torsional angle profile through steric hin-
drance between H6 and sugar hydrogens and, thereby, the
positioning and photophysical properties of the C5-fluoro-
phore (see Figure S14 in the Supporting Information).
Figure 4. Fluorescence intensity of ON5–ON16 in the absence (SSPs) or
presence of complementary DNA or mismatched DNA targets. Panels
depict ONs modified with monomer X (upper), monomer Y (middle), or
monomer Z (lower). Hybridization-induced increases and discrimination
factors (Im/Imm), defined as the fluorescence intensity of duplexes with
complementary DNA divided by the intensity of SSPs or duplexes with
mismatched DNA, respectively, are listed above corresponding histo-
grams. Intensity recorded at lem =402 nm at T=58C, 1.0 mm concentra-
tion of each strand.
duplexes than in matched duplexes (Table 2). It is particular-
ly noteworthy that larger relative differences in extinction
coefficients between matched and mismatched duplexes are
observed with ON9–ON16 (LNA and a-l-LNA monomers
Y and Z) than with ON5–ON8. This demonstrates that a)
decreased quantum yields in tandem with lower extinction
coefficients of mismatched duplexes bring about the SNP
discrimination of monomers X–Z, and b) conformational re-
striction of the furanose skeleton translates into altered
emission output of polarity-sensitive fluorophores conjugat-
ed to the C5 position of pyrimidines.
Conclusion
Hybridization to RNA targets: Motivated by these results
and the importance of fluorescent probes in the elucidation
of biological functions of RNA,[23] we set out to study the
physical properties of duplexes between probes in the repre-
sentative CBC- and TBT-sequence contexts and comple-
The study provides important insight into the fluorescence
properties of SNP-typing probes utilizing 5-[3-(1-pyrenecar-
boxamido)propynyl]pyrimidines as base discriminating fluo-
rescent monomers. First, efficient optical discrimination of
3162
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ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Chem. Eur. J. 2011, 17, 3157 – 3165

SNP Discrimination with LNA
FULL PAPER
74%) as a light yellow solid. Rf =0.5 (5% MeOH in CH2Cl2, v/v); ESI-
HRMS: m/z: 862.2778 ([M+Na]+, C51H41N3O9+Na, calcd 862.2735);
1H NMR ([D6]DMSO): d=1.71 (s, 1H, ex, NH), 9.95 (t, 1H, ex, J=
5.5 Hz, NHCH2), 8.52–8.54 (d, 1H, J=9.5 Hz, Ar), 8.21–8.36 (m, 6H,
Ar), 8.09–8.13 (m, 2H, Ar), 7.84 (s, 1H, H6), 7.23–7.46 (m, 9H, Ar), 6.90
(d, 4H, J=9.0 Hz, Ar), 5.73 (d, 1H, ex, J=4.5 Hz, 3’-OH), 5.45 (s, 1H,
H1’), 4.20–4.34 (m, 3H, H2’, CH2NH), 4.07 (d, 1H, J=4.5 Hz, H3’), 3.82–
3.83 (d, 1H, J=8.0 Hz, H5’’), 3.80–3.81 (d, 1H, J=8.0 Hz, H5’’), 3.72 (s,
6H, 2ꢃOCH3), 3.57–3.59 (d, 1H, J=11.0 Hz, H5’), 3.29–3.31 ppm (d,
1H, J=11.0 Hz, H5’); 13C NMR ([D6]DMSO): d=168.5, 161.8, 158.12,
158.07, 149.0, 144.7, 141.6 (C6), 135.5, 134.9, 131.7, 131.0, 130.7, 130.1,
129.9 (Ar), 129.6 (Ar), 128.3 (Ar), 128.1 (Ar), 127.90 (Ar), 127.89, 127.5
(Ar), 127.1 (Ar), 126.7 (Ar), 126.5 (Ar), 125.8 (Ar), 125.2 (Ar), 124.6
(Ar), 124.3 (Ar), 123.7, 123.6, 113.3 (Ar), 113.2 (Ar), 97.8, 89.4, 87.6, 87.0
(C1’), 85.6, 78.8 (C2’), 74.6, 71.4 (C5’), 69.7 (C3’), 59.1 (C5’’), 54.98
(OCH3), 54.97 (OCH3), 29.5 ppm (CH2NH).
SNPs by ONs modified with known BDF monomer X is
demonstrated to necessitate nearby guanine moieties.
Second, conformational restriction of the furanose skeleton
(monomers Y and Z) translates into altered emission output
of polarity-sensitive fluorophores appended to the C5 posi-
tion of pyrimidines, which reflects changes in quantum
yields and extinction coefficients of the fluorophore. Ac-
cordingly, probes modified with LNA and a-l-LNA mono-
mers Y and Z display more beneficial SNP-typing character-
istics. We envision that the use of ONs modified with LNA/
a-l-LNA monomers conjugated to polarity-sensitive fluoro-
phores via short rigid linkers is a promising strategy toward
development of probes for sequence-unrestricted SNP-
typing.
(1S,3R,4S,7R)-1-(4,4’-Dimethoxytrityloxymethyl)-7-hydroxy-3-[5-{3-(1-
pyrenecarboxamido)propynyl}uracil-1-yl]-2,5-dioxabicyclo
(2Z): Nucleoside 1Z (0.50 g, 0.73 mmol), [Pd(PPh3)4] (90 mg,
0.07 mmol), CuI (30 mg, 0.14 mmol), N-(prop-2-ynyl)pyrene-1-carbox-
amide[8d] (0.28 g, 1.00 mmol), and Et3N (0.40 mL, 2.84 mmol) in anhy-
ACHTUNGTREN[NGNU 2.2.1]heptane
AHCTUNGTRENNUNG
Experimental Section
AHCTUNGTRENNUNG
drous DMF (10 mL) were allowed to react, worked up, and purified as
described for 2Y to provide nucleoside 2Z (0.47 g, 79%) as a light
yellow solid material. Rf =0.5 (5% MeOH in CH2Cl2, v/v); ESI-HRMS:
m/z: 862.2756 ([M+Na]+, C52H41N3O9 Na+, calcd 862.2735); 1H NMR
([D6]DMSO): d=11.77 (s, 1H, ex, NH), 9.23 (t, 1H, ex, J=5.5 Hz,
NHCH2), 8.54–8.55 (d, 1H, J=9.5 Hz, Ar), 8.19–8.35 (m, 6H, Ar), 8.10–
8.14 (m, 2H, Ar), 7.99 (s, 1H, H6), 7.13–7.40 (m, 9H, Ar), 6.88 (d, 4H,
J=9.0 Hz, Ar), 5.98 (s, 1H, H1’), 5.95 (d, 1H, ex, J=4.5 Hz, 3’-OH), 4.49
(d, 2H, J=5.5 Hz, CH2NH), 4.43 (d, 1H, J=4.5 Hz, H3’), 4.28 (s, 1H,
H2’), 4.03–4.05 (d, 1H, J=8.5 Hz, H5’’), 3.95–3.97 (d, 1H, J=8.5 Hz,
H5’’), 3.69 (s, 6H, 2ꢃCH3O), 3.32 ppm (s, 2H, H5’); 13C NMR
([D6]DMSO): d=168.6, 161.7, 158.1, 149.3, 144.7, 143.0 (C6), 135.2,
135.1, 131.7, 131.0, 130.7, 130.1, 129.79 (Ar), 129.67 (Ar), 128.3 (Ar),
128.1 (Ar), 127.9 (Ar), 127.8 (Ar), 127.5 (Ar), 127.1 (Ar), 126.6 (Ar),
126.5 (Ar), 125.8 (Ar), 125.6 (Ar), 125.2 (Ar), 124.5 (Ar), 124.3 (Ar),
123.8, 123.6, 113.2 (Ar), 97.1, 89.5, 89.4, 87.1 (C1’), 85.3, 78.7 (C2’), 74.5,
72.8 (C3’), 72.4 (C5’’), 59.8 (C5’), 54.9 (CH3O), 29.6 ppm (CH2NH).
General: All reagents and solvents were of analytical grade and obtained
from commercial suppliers and used without further purification. Petrole-
um ether of the distillation range 60–808C was used. Anhydrous di-
chloromethane, 1,2-dichloroethane, and N,N-diisopropylethylamine
(DIPEA) were dried through storage over activated 4 ꢂ molecular
sieves. Water content of the anhydrous solvents was checked by a Karl–
Fischer apparatus. Reactions were conducted under an atmosphere of
argon whenever anhydrous solvents were used. All reactions were moni-
tored by thin-layer chromatography (TLC) using silica gel coated plates
with fluorescence indicator (SiO2-60, F-254) which were visualized a)
under UV light or, b) by dipping in 5% conc. sulfuric acid 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 moder-
ate pressure (pressure ball). Silica gel columns were built with an initial
starting eluent containing 1% (v/v) of pyridine. Evaporation of solvents
was carried out under reduced pressure at temperatures below 508C.
After column chromatography, appropriate fractions were pooled, evapo-
rated and dried at high vacuum for at least 12 h to give the obtained
products in high purity (>95%) which was ascertained by 1D NMR tech-
niques. Chemical shifts of 1H NMR (500 MHz), 13C NMR (125 MHz),
and 31P NMR (121.5 MHz) spectra are reported in parts per million
(ppm) relative to deuterated solvent or other internal standards (80%
phosphoric acid for 31P NMR, respectively). Exchangeable (ex) protons
were detected by disappearance of peaks on D2O addition. Assignments
of NMR spectra are based on 2D spectra (COSY, HSQC) and DEPT
spectra. Quaternary carbons were not assigned but verified from HSQC
and DEPT spectra (absence of signals). The carbon atom of C4’ substitu-
ents is numbered C5’’. Similar conventions apply for the corresponding
hydrogen atoms. Assignments of 1H NMR signals of H5ꢁ and H5’ꢁ and
the corresponding 13C NMR signals are interchangeable. ESI-HRMS
spectra of compounds dissolved in a saturated solution of NaCl in
CH3CN with PEG as an internal calibrant, were recorded on a Quadro-
pole Time-Of-Flight tandem Mass Spectrometer (Q-TOF Premiere).
(1R,3R,4R,7S)-7-[2-Cyanoethoxy(diisopropylamino)phosphinoxy]-1-(4,4’-
dimethoxytrityloxymethyl)-3-[5-{3-(1-pyrenecarboxamido)propynyl}ura-
cil-1-yl]-2,5-dioxabicycloACTHUNRTGNEUNG[2.2.1]heptane (3Y): Alcohol 2Y (0.25 g,
0.29 mmol) was coevaporated with anhydrous 1,2-dichloroethane (2ꢃ
10 mL) and dissolved in anhydrous CH2Cl2. To this was added DIPEA
(0.19 mL, 1.50 mmol), and 2-cyanoethyldiisopropylchlorophosporamidite
(PCl reagent) (0.09 mL, 0.38 mmol) and the reaction mixture was stirred
at room temperature for 2 h. The reaction mixture was diluted with
CH2Cl2 (25 mL), washed with 5% aq. NaHCO3 (2ꢃ10 mL), and the com-
bined aqueous phase back-extracted with CH2Cl2 (2ꢃ10 mL). The com-
bined organic phase was dried (Na2SO4), evaporated to dryness, and the
resulting crude residue purified by column chromatography (0–2%
MeOH/CH2Cl2, v/v) to provide phosphoramidite 3Y (190 mg, 61%) as a
white foam. Rf =0.5 (2% MeOH in CH2Cl2, v/v); ESI-HRMS: m/z:
1062.3807 ([M+Na]+, C60H58N5O10P·Na+, calcd 1062.3814); 31P NMR
(CDCl3, 121.5 MHz): d=149.7, 149.2 ppm.
(1S,3R,4S,7R)-7-[2-Cyanoethoxy(diisopropylamino)phosphinoxy]-1-(4,4’-
dimethoxytrityloxymethyl)-3-[5-{3-(1-pyrenecarboxamido)propynyl}ura-
cil-1-yl]-2,5-dioxabicycloACTHNUGRTENUNG[2.2.1]heptane (3Z): Nucleoside 2Z (0.44 g,
(1R,3R,4R,7S)-1-(4,4’-Dimethoxytrityloxymethyl)-7-hydroxy-3-[5-{3-(1-
pyrenecarboxamido)propynyl}uracil-1-yl]-2,5-dioxabicyclo
ACHTUNGTREN[NGNU 2.2.1]heptane
(2Y): Nucleoside 1Y (0.50 g, 0.73 mmol), [Pd(PPh3)4] (90 mg,
AHCTUNGTRENNUNG
0.52 mmol), DIPEA (0.46 mL, 2.6 mmol), and 2-cyanoethyldiisopropyl-
chlorophosporamidite (0.18 mL, 0.78 mmol) in anhydrous CH2Cl2
(10 mL) were allowed to react, worked up, and purified as described for
3Y to furnish phosphoramidite 3Z (0.41 g, 75%) as a white foam. Rf =
0.5 (2% MeOH in CH2Cl2, v/v); ESI-HRMS: m/z: 1062.3790 ([M+Na]+,
C60H58N5O10P·Na+, calcd 1062.3814); 31P NMR (CDCl3, 121.5 MHz): d=
150.3, 149.8 ppm.
0.07 mmol), CuI (30 mg, 0.14 mmol), and N-(prop-2-ynyl)pyrene-1-car-
boxamide[8d] (0.28 g, 1.00 mmol) were added to anhydrous DMF (10 mL)
and the resulting mixture was degassed and placed under argon. To this
was added Et3N (0.40 mL, 2.84 mmol) and the reaction mixture was
stirred at room temperature for 12 h whereupon solvents were evaporat-
ed off. The resulting residue was taken up in EtOAc (100 mL) and se-
quentially washed with brine (2ꢃ50 mL) and sat. aq. NaHCO3 (50 mL).
The combined aqueous phase was back-extracted with EtOAc (100 mL),
and the combined organic phase was dried (Na2SO4), evaporated to dry-
ness and the resulting crude residue purified by column chromatography
(0–5% MeOH in CH2Cl2 (v/v)) to afford the nucleoside 2Y (0.45 g,
Oligonucleotide synthesis: Oligonucleotides (ONs) were synthesized on a
0.2 mmol scale using an Expedite 8909 Synthesizer and succinyl-linked
LCAA-CPG (long-chain alkyl amine controlled pore glass) columns with
a pore size of 500 ꢂ. Standard procedures were used, that is, trichloro-
Chem. Eur. J. 2011, 17, 3157 – 3165
ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chemeurj.org
3163

P. J. Hrdlicka et al.
acetic acid in CH2Cl2 as detritylation reagent; 0.25m 4,5-dicyanoimidazole
(DCI) in anhydrous CH3CN as activator; acetic anhydride in THF as cap
A solution; 1-methylimidazole in THF/pyridine (8:1, v/v) as cap B solu-
tion, and 0.02m iodine in H2O/pyridine/THF as the oxidizing solution. In-
corporation of monomers X–Z into ONs was accomplished by extended
coupling (15 min) of the corresponding phosphoramidites 3X–Z with
DCI as the activator, which resulted in coupling yields above 98%.
Cleavage from the solid support and removal of protecting groups was
accomplished by treatment with concentrated aqueous ammonia (558C,
24 h). Purification of all modified ONs was performed to minimum 80%
purity using either of two methods: a) overall synthesis yield >80%:
cleavage of DMT using 80% aq. AcOH, followed by precipitation from
acetone (ꢀ188C for 12–16 h) and washing with acetone, or b) overall syn-
thesis <80%: purification of ONs by RP-HPLC using a gradient of
0.05m triethyl ammonium acetate in water and 25% water in acetonitrile
(see Table S1 in the Supporting Information), followed by detritylation
and precipitation as outlined above.
plexes with DNA/RNA targets were determined according to Equa-
tion (1),
FFðONÞ ¼ FFðPBAÞ ꢃ ½AðONÞ=A344 ðONÞꢄ ꢃ ½1=aðPBAÞꢄ
ð1Þ
2
2 ½17ꢄ
ꢃ ½nðH2OÞ =nðMeOHÞ ꢄ
where FFACTHUNRTGNEUNG(PBA) is the cross-calibrated value for the fluorescence quan-
tum yield of PBA in MeOH, A(ON) is the area of the fluorescence emis-
sion spectrum of the sample from 360 to 600 nm, A344 (ON) is the absorb-
ance of the sample at the excitation wavelength (344 nm), a
slope of the fluorescence emission versus A344 (ON) calibration curve for
PBA and n(H2O) and n(MeOH) are the refractive indexes of water
ACHTUNGTRENUN(NG PBA) is the
A
ACHTUNGTRENNUNG
(1.3328) and methanol (1.3288), respectively. The reported quantum
yields are an average of at least two measurements within (ꢂ10)%, al-
though low quantum yields (FF <10%) may be associated with consider-
ably larger error.
Fluorescence excitation spectra were recorded using the same buffer and
concentrations as for the thermal denaturation studies at T=58C. 402 nm
was used as the emission wavalength and excitation intensity was scanned
from 300 to 400 nm.
Purification of crude ONs was performed on a Varian Prostar HPLC
system equipped with an XTerra MS C18 column (10 mm, 7.8ꢃ150 mm)
using the representative gradient protocol depicted in Table S1. The com-
position of all synthesized ONs was verified by MALDI MS analysis (see
Table S2) recorded in positive ion mode on a Quadropole Time-Of-
Flight tandem Mass Spectrometer (Q-TOF Premiere) equipped with a
MALDI source (Waters Micromass LTD., U.K.) using 2,5-dihydroxyben-
zoic acid as a matrix and PEG as an internal standard. Purity (>80%)
was verified by RP-HPLC.
Acknowledgements
We appreciate financial support from Idaho NSF EPSCoR, the BANTech
Center at the University of Idaho, and a scholarship from the College of
Graduate Students, University of Idaho (M.E.Ø.). We thank the EBI
Murdock Mass Spectrometry Center for mass spectrometric analyses.
F.M.Y. thanks IBEST and BANTech at the University of Idaho for com-
puting resources.
Protocol for thermal denaturation studies: Concentrations of unmodified
ONs were estimated using the following extinctions coefficients for DNA
(OD/mmol): G (12.01), A (15.20), T (8.40), C (7.05); for RNA (OD/
mmol): G (13.70), A (15.40), U (10.00), C (9.00). Concentrations of modi-
fied ONs (ON5–ON16) were determined by titration with complementa-
ry DNA; a progressive increase in steady-state fluorescence emission in-
tensity was observed until a plateau was reached, at which point a 1:1
stoichiometry was assumed. This approach was crossvalidated by compar-
ison with conventional methods for concentration determination of
pyrene-functionalized ONs which assume an e260 of 22.4 OD/mmol for the
pyrene moiety.[10b] The two methods gave concentration determinations
within (ꢂ10)%. ONs (1.0 nmol of each strand, 1 mm) were thoroughly
mixed, denatured by heating and subsequently cooled to the starting tem-
perature of the experiment. Quartz optical cells with a pathlength of
1.0 cm were used. Thermal denaturation temperatures (Tm values [8C])
were measured on a Cary 100 UV/VIS spectrophotometer equipped with
12-cell Peltier temperature controller and determined as the maximum of
the first derivative of the thermal denaturation curve (A260 vs. T) record-
ed in medium salt buffer (Tm buffer: 110 mm NaCl, 0.1 mm EDTA, pH
adjusted with 10 mm Na2HPO4/NaH2PO4). The temperature of the dena-
turation experiments ranged from at least 158C below Tm to 208C above
Tm (although not below 18C). A temperature ramp of 0.58Cminꢀ1 was
used in all experiments. Reported thermal denaturation temperatures are
an average of at least two experiments within (ꢂ1.0)8C.
[1] S. Kim, A. Misra, Annu. Rev. Biomed. Eng. 2007, 9, 289–320.
[2] C. Ahlborn, K. Siegmund, C. Richert, J. Am. Chem. Soc. 2007, 129,
15218–15232.
[3] a) N. Venkatesan, Y. J. Seo, B. H. Kim, Chem. Soc. Rev. 2008, 37,
648–663; b) K. Wang, Z. Tang, C. J. Yang, Y. Kim, X. Fang, W. Li,
Y. Wu, C. D. Medley, Z. Cao, J. Li, P. Colon, H. Lin, W. Tan, Angew.
Chem. 2009, 121, 870–885; Angew. Chem. Int. Ed. 2009, 48, 856–
870.
[4] a) D. M. Kolpashchikov, Chem. Rev. 2010, 110, 4709–4723; b) P. L.
Paris, J. M. Langenhan, E. T. Kool, Nucleic Acids Res. 1998, 26,
3789–3793; c) A. A. Marti, S. Jockusch, N. Stevens, J. Ju, N. J. Turro,
Acc. Chem. Res. 2007, 40, 402–409; d) T. Umemoto, P. J. Hrdlicka,
B. R. Babu, J. Wengel, ChemBioChem 2007, 8, 2240–2248.
[5] a) A. P. Silverman, E. T. Kool, Chem. Rev. 2006, 106, 3775–3789;
b) T. N. Grossmann, O. Seitz, Chem. Eur. J. 2009, 15, 6723–6730.
[6] a) U. B. Christensen, E. B. Pedersen, Helv. Chim. Acta 2003, 86,
2090–2097; b) O. Kçhler, D. Venkatrao, D. V. Jarikote, O. Seitz,
ChemBioChem 2005, 6, 69–77; c) T. S. Kumar, J. Wengel, P. J.
Hrdlicka, ChemBioChem 2007, 8, 1122–1125.
[7] a) A. A. Gorodetsky, M. C. Buzzeo, J. K. Barton, Bioconjugate
Chem. 2008, 19, 2285–2296; b) L. Valis, N. Amann, H. A. Wagen-
knecht, Org. Biomol. Chem. 2005, 3, 36–38; c) E. Mayer-Enthart,
H. A. Wagenknecht, Angew. Chem. 2006, 118, 3451–3453; Angew.
Chem. Int. Ed. 2006, 45, 3372–3375; d) C. Wanninger-Weiss, L.
Valis, H. A. Wagenknecht, Bioorg. Med. Chem. 2008, 16, 100–106.
[8] a) A. Okamoto, Y. Saito, I. Saito, J. Photochem. Photobiol. C 2005,
6, 108–122; b) D. W. Dodd, R. H. E. Hudson, Mini-Rev. Org. Chem.
2009, 6, 378–391; c) K. Yamana, H. Zako, K. Asazuma, R. Iwase, H.
Nakano, A. Murakami, Angew. Chem. 2001, 113, 1138–1140;
Angew. Chem. Int. Ed. 2001, 40, 1104–1106; d) A. Okamoto, K. Ka-
natani, I. Saito, J. Am. Chem. Soc. 2004, 126, 4820–4827; e) G. T.
Hwang, Y. J. Seo, S. J. Kim, B. H. Kim, Tetrahedron Lett. 2004, 45,
3543–3546; f) R. H. E. Hudson, A. Ghorbani-Choghamarani, Org.
Biomol. Chem. 2007, 5, 1845–1848; g) M. E. Østergaard, D. C.
Guenther, P. Kumar, B. Baral, A. J. Paszczynski, P. K. Sharma, P. J.
Hrdlicka, Chem. Commun. 2010, 46, 4929–4931; h) S. S. Bag, R.
Protocol for fluorescence studies: Steady state fluorescence emission
spectra were recorded using a Cary Eclipse fluorimeter using the same
buffers and ON concentrations (1.0 mm) as in thermal denaturation stud-
ies. Fluorescence emission spectra of single stranded probes (SSP) and
corresponding duplexes with complementary or mismatched targets were
measured at 58C to ensure maximal hybridization. Deoxygenation was
deliberately not applied to the samples since the scope of the work was
to determine fluorescence under aerated condition prevailing in bioas-
says. Solutions were heated to 808C over 10 min, cooled to 58C over
15 min, and equilibrated at this temperature for more than 5 min. Steady-
state fluorescence emission spectra (360–600 nm range) were obtained as
an average of five scans using an excitation wavelength of 344 nm, excita-
tion slit 5.0 nm, emission slit 5.0 nm and a scan speed of 600 nmminꢀ1
.
The fluorescence quantum yield FF of pyrenebutanoic acid (PBA) in
MeOH in this experimental setting was measured to be 0.069 relative to
9,10-diphenylanthracene in cyclohexene (FF =0.95),[17] which is in excel-
lent agreement with the reported value of 0.065.[18] Emission quantum
yields FF (ON) of single-stranded ON9–ON12 and the corresponding du-
3164
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ꢀ 2011 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Chem. Eur. J. 2011, 17, 3157 – 3165

SNP Discrimination with LNA
FULL PAPER
Kundu, K. Matsumoto, Y. Saito, I. Saito, Bioorg. Med. Chem. Lett.
2010, 20, 3227–3230; i) Y. Shinohara, K. Matsumoto, K. Kugenuma,
T. Morii, Y. Saito, I. Saito, Bioorg. Med. Chem. Lett. 2010, 20, 2817–
2820.
Petersen, A. E. Hꢄkansson, J. Wengel, J. P. Jacobsen, J. Am. Chem.
Soc. 2001, 123, 7431–7432.
[14] W. Saenger, Principles of Nucleic Acid Structure, Springer, Berlin,
1984.
[9] A. Okamoto, K. Tainaka, Y. Ochi, K. Kanatani, I. Saito, Mol. Bio-
Syst. 2006, 2, 122–126.
[15] a) V. A. Korshun, D. A. Stetsenko, M. J. Gait, J. Chem. Soc. Perkin
Trans. 1 2002, 1092–1104; b) C. Dohno, I. Saito, ChemBioChem
2005, 6, 1075–1081.
[16] Y. You, B. G. Moreira, M. A. Behlke, R. Owczarzy, Nucleic Acids
Res. 2006, 34, e60.
[17] J. V. Morris, M. A. Mahaney, J. R. Huber, J. Phys. Chem. 1976, 80,
969–974.
[18] T. L. Netzel, K. Nafisi, J. Headrick, B. E. Eaton, J. Phys. Chem.
1995, 99, 17948–17955.
[19] G. Dougherty, J. R. Pilbrow, Int. J. Biochem. 1984, 16, 1179–1192.
[20] a) P. J. Hrdlicka, B. R. Babu, M. D. Sørensen, N. Harrit, J. Wengel, J.
Am. Chem. Soc. 2005, 127, 13293–13299; b) D. Honcharenko, C.
Zhou, J. Chattopadhyaya, J. Org. Chem. 2008, 73, 2829–2842;
c) I. V. Astakhova, V. A. Korshun, J. Wengel, Chem. Eur. J. 2008, 14,
11010–11026.
[21] M. Nakamura, Y. Fukunaga, K. Sasa, Y. Ohtoshi, K. Kanaori, H.
Hayashi, H. Nakano, K. Yamana, Nucleic Acids Res. 2005, 33, 5887–
5895.
[10] a) T. S. Kumar, A. S. Madsen, M. E. Østergaard, J. Wengel, P. J.
Hrdlicka, J. Org. Chem. 2008, 73, 7060–7066; b) T. S. Kumar, A. S.
Madsen, M. E. Østergaard, S. P. Sau, J. Wengel, P. J. Hrdlicka, J.
Org. Chem. 2009, 74, 1070–1081; c) M. E. Østergaard, P. Kumar, B.
Baral, D. J. Raible, T. S. Kumar, B. A. Anderson, D. C. Guenther, L.
Deobald, A. J. Paszczynski, P. K. Sharma, P. J. Hrdlicka, ChemBio-
Chem 2009, 10, 2740–2743; d) S. P. Sau, P. Kumar, B. A. Anderson,
M. E. Østergaard, L. Deobald, A. Paszczynski, P. K. Sharma, P. J.
Hrdlicka, Chem. Commun. 2009, 6756–6758; e) S. P. Sau, T. S.
Kumar, P. J. Hrdlicka, Org. Biomol. Chem. 2010, 8, 2028–2036;
f) M. E. Østergaard, P. Cheguru, M. R. Papasani, R. A. Hill, P. J.
Hrdlicka, J. Am. Chem. Soc. 2010, 132, 14221–14228.
[11] a) A. A. Koshkin, S. K. Singh, P. Nielsen, V. K. Rajwanshi, R.
Kumar, M. Meldgaard, C. E. Olsen, J. Wengel, Tetrahedron 1998, 54,
3607–3630; b) S. Obika, T. Uneda, T. Sugimoto, D. Nanbu, T.
Minami, T. Doi, T. Imanishi, Bioorg. Med. Chem. 2001, 9, 1001–
1011; c) H. Kaur, B. R. Babu, S. Maiti, Chem. Rev. 2007, 107, 4672–
4697.
[22] M. Manoharan, K. L. Tivel, M. Zhao, K. Nafisi, T. L. Netzel, J. Phys.
Chem. 1995, 99, 17461–17472.
[12] M. D. Sørensen, L. Kværnø, T. Bryld, A. E. Hꢄkansson, B. Verbeure,
G. Gaubert, P. Herdewijn, J. Wengel, J. Am. Chem. Soc. 2002, 124,
2164–2176.
[23] S. Tyagi, Nat. Methods 2009, 6, 331–338.
[24] V. Marin, H. F. Hansen, T. Koch, B. A. Armitage, J. Biomol. Struct.
Dyn. 2004, 21, 841–850.
[13] a) M. Petersen, C. B. Nielsen, K. E. Nielsen, G. A. Jensen, K. Bon-
densgaard, S. K. Singh, V. K. Rajwanshi, A. A. Koshkin, B. M. Dahl,
J. Wengel, J. P. Jacobsen, J. Mol. Recognit. 2000, 13, 44–53; b) M.
Received: July 23, 2010
Revised: November 16, 2010
Published online: February 15, 2011
Chem. Eur. J. 2011, 17, 3157 – 3165
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3165