Recognition of double-stranded DNA using energetically activated duplexes with interstrand zippers of 1-, 2- or 4-pyrenyl-functionalized O2′-alkylated RNA monomers

Article abstract of DOI:10.1039/c4ob01183j

Despite advances with triplex-forming oligonucleotides, peptide nucleic acids, polyamides and-more recently-engineered proteins, there remains an urgent need for synthetic ligands that enable specific recognition of double-stranded (ds) DNA to accelerate studies aiming at detecting, regulating and modifying genes. Invaders, i.e., energetically activated DNA duplexes with interstrand zipper arrangements of intercalator-functionalized nucleotides, are emerging as an attractive approach toward this goal. Here, we characterize and compare Invaders based on 1-, 2- and 4-pyrenyl-functionalized O2′-alkylated uridine monomers X-Z by means of thermal denaturation experiments, optical spectroscopy, force-field simulations and recognition experiments using DNA hairpins as model targets. We demonstrate that Invaders with +1 interstrand zippers of X or Y monomers efficiently recognize mixed-sequence DNA hairpins with single nucleotide fidelity. Intercalator-mediated unwinding and activation of the double-stranded probe, coupled with extraordinary stabilization of probe-target duplexes (ΔT_m/modification up to +14.0 °C), provides the driving force for dsDNA recognition. In contrast, Z-modified Invaders show much lower dsDNA recognition efficiency. Thus, even very conservative changes in the chemical makeup of the intercalator-functionalized nucleotides used to activate Invader duplexes, affects dsDNA-recognition efficiency of the probes, which highlights the importance of systematic structure-property studies. The insight from this study will guide future design of Invaders for applications in molecular biology and nucleic acid diagnostics.

Full text of DOI:10.1039/c4ob01183j

Organic &
Biomolecular Chemistry
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Recognition of double-stranded DNA using
energetically activated duplexes with interstrand
zippers of 1-, 2- or 4-pyrenyl-functionalized
O2’-alkylated RNA monomers†
Cite this: Org. Biomol. Chem., 2014,
12, 7758
Saswata Karmakar,a Andreas S. Madsen,b Dale C. Guenther,a Bradley C. Gibbonsa,c
and Patrick J. Hrdlicka*a
Despite advances with triplex-forming oligonucleotides, peptide nucleic acids, polyamides and – more
recently – engineered proteins, there remains an urgent need for synthetic ligands that enable specific
recognition of double-stranded (ds) DNA to accelerate studies aiming at detecting, regulating and modify-
ing genes. Invaders, i.e., energetically activated DNA duplexes with interstrand zipper arrangements of
intercalator-functionalized nucleotides, are emerging as an attractive approach toward this goal. Here, we
characterize and compare Invaders based on 1-, 2- and 4-pyrenyl-functionalized O2’-alkylated uridine
monomers X–Z by means of thermal denaturation experiments, optical spectroscopy, force-field simu-
lations and recognition experiments using DNA hairpins as model targets. We demonstrate that Invaders
with +1 interstrand zippers of X or Y monomers efficiently recognize mixed-sequence DNA hairpins with
single nucleotide fidelity. Intercalator-mediated unwinding and activation of the double-stranded probe,
coupled with extraordinary stabilization of probe–target duplexes (ΔTm/modification up to +14.0 °C), pro-
vides the driving force for dsDNA recognition. In contrast, Z-modified Invaders show much lower dsDNA
recognition efficiency. Thus, even very conservative changes in the chemical makeup of the intercalator-
functionalized nucleotides used to activate Invader duplexes, affects dsDNA-recognition efficiency of the
probes, which highlights the importance of systematic structure–property studies. The insight from this
study will guide future design of Invaders for applications in molecular biology and nucleic acid
diagnostics.
Received 9th June 2014,
Accepted 4th August 2014
DOI: 10.1039/c4ob01183j
www.rsc.org/obc
DNA (dsDNA) has proven very challenging as the Watson–Crick
base pairs are buried deeply within the duplex core and not
Introduction
The right-handed DNA helix is one of the most fundamental readily accessible to exogenous agents. The most established
structures in Nature due to its role as the carrier of genetic dsDNA-targeting agents, i.e., pyrrole–imidazole polyamides,2
information. The two strands comprising the helix are held engineered proteins3 and triplex-forming oligonucleotides4 or
together through π–π stacking interactions between neighbor- peptide nucleic acids (PNAs),5 accordingly recognize chemical
ing nucleobases and hydrogen bonding between base pairs.1 features that are accessible via one of the grooves instead.6
Development of synthetic ligands that are capable of decoding While these strategies are attractive, they are not without limit-
the sequence information contained within double-stranded ations. For example, PNA-based approaches normally require
low salinity conditions, while triplex-based approaches require
targets with polypurine tracts. Approaches with more relaxed
sequence requirements have been developed but they still
aDepartment of Chemistry, University of Idaho, Moscow, ID 83844, USA.
E-mail: hrdlicka@uidaho.edu; Fax: (+1) 208 885 6173; Tel: (+1) 208 885 0108
bDepartment of Chemistry, Technical University of Denmark, Kemitorvet 207,
require the presence of purine stretches, a circumstance that
may not be met at a target of interest.7 Other strategies allow
Kgs. Lyngby, DK-2800, Denmark
cDepartment of Chemistry, Brigham Young University-Idaho, ID 83440, USA
†Electronic supplementary information (ESI) available: General experimental
recognition of mixed-sequence target regions that are un-
usually accessible to exogenous agents, such as AT-rich cruci-
forms or transcription bubbles.8
Synthetic ligands that recognize mixed-sequence B-DNA via
duplex invasion are attractive due to the predictability of
Watson–Crick base-pairing rules. However, these approaches
section; MS-data for modified ONs; representative thermal denaturation curves;
additional denaturation and thermodynamic data; additional absorption and
steady-state fluorescence emission spectra; complete data set of structural para-
meters from modeling studies; lowest energy representations of Type B struc-
tures; NMR spectra for nucleosides 2Y–4Z. See DOI: 10.1039/c4ob01183j
7758 | Org. Biomol. Chem., 2014, 12, 7758–7773
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Fig. 1 Illustration of the Invader approach for recognition of mixed-sequence DNA and structures of monomers used herein. Droplets denote
pyrene moieties.
must overcome a steep energetic penalty since pre-existing DNA in male bovine kidney cells at non-denaturing
base pairs of B-DNA must be broken prior to probe binding. conditions.13
Pseudocomplementary DNA (pcDNA), i.e., DNA duplexes fea-
We originally used 2′-N-(pyren-1-yl)methyl-2′-amino-α-L-LNA
turing modified nucleobases that form weak base pairs with (locked nucleic acid) nucleotides as the key activating com-
each other, while allowing hybridization to complementary ponents of Invader probes,15 but recently discovered that they
DNA, recognize mixed-sequence target regions at B-DNA can be replaced by 2′-O-(pyren-1-yl)methyl-ribonucleotides
termini.9 This strategy has been successfully extended to (Fig. 1).16 The resulting probes display similar dsDNA-recog-
pcPNA, which recognize internal regions of mixed-sequence nition efficiency and are much easier to synthesize.17,18 Identi-
dsDNA.10 However, the self-inhibitory effects observed at high fication of these simple building blocks allowed us to initiate
pcPNA concentrations and the requirement for low salinity, systematic structure–property relationship studies with the
pose potential limitations for pcPNA-mediated strand invasion goal of gaining additional insights into the structural determi-
in biological media.11 γ-PNA, i.e., single-stranded probes that nants governing Invader recognition efficiency. For example,
are fully modified with conformationally pre-organized PNA we demonstrated that all four canonical 2′-O-(pyren-1-yl)-
building blocks, are another interesting class of dsDNA-target- methyl-RNA monomers can be used to construct dsDNA-
ing probes capable of recognizing mixed-sequence dsDNA binding Invader probes. However, probes using the pyrimidine
targets. However, they too require non-physiological salinity monomers are particularly efficient.19
for optimal strand invasion.12 In summary, oligonucleotide-
based probes that recognize mixed-sequence dsDNA targets at orientation between the pyrene intercalator and sugar skeleton
physiologic conditions remain largely elusive. has any impact on Invader-mediated recognition of mixed-
In the present study, we wanted to determine if the relative
We have recently introduced Invaders as an alternative sequence dsDNA. Toward this end, the corresponding 2- and
strategy toward mixed-sequence dsDNA recognition.13 These 4-pyrenyl functionalized uridines were synthesized and incor-
double-stranded probes are activated for dsDNA recognition porated into oligodeoxyribonucleotides (ONs), which were
through modification with +1 interstrand zippers of inter- then characterized with respect to thermal denaturation, ther-
calator-functionalized nucleotides (for an illustration, see modynamic, UV-Vis absorption and fluorescence properties
Fig. 1; for a formal definition of the zipper nomenclature, (Fig. 1). This was then followed by biophysical characterization
see the Experimental section). This structural motif results in of double-stranded probes with different interstrand arrange-
a locally perturbed and destabilized region in the probe ments of these monomers and recognition experiments using
duplex since the motif represents a violation of the ‘nearest DNA hairpins as model targets.
neighbor exclusion principle’, which states that intercalators,
at most, bind to every second base pair of a DNA duplex due to
limitations in local helix expandability.14 On the other
hand, the two strands comprising an Invader probe display
exceptionally strong affinity toward complementary single-
stranded DNA (cDNA), since duplex formation results in
Results and discussion
Synthesis of O2′-pyrene-functionalized uridine
phosphoramidites
strongly stabilizing pyrene–nucleobase stacking interactions
(Fig. 1). In a key proof-of-concept study, we harnessed the
energy difference between reactants (i.e., Invader probes
and target duplexes) and products (i.e., probe–target duplexes)
to drive recognition of chromosomal DYZ-1 satellite
Phosphoramidites 4Y and 4Z were prepared in a similar
manner as the 2′-O-(pyren-1-yl)methyl-uridine analogue 4X.17b
Thus, treatment of O2,O2′-anhydrouridine 120 with tris(pyren-
2-yl)methyl borate or tris(pyren-4-yl)methyl borate – generated
in situ21 via addition of 2-pyrenemethanol22 or 4-pyrenemetha-
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Scheme 1 Synthesis of O2’-pyrene-functionalized uridine phosphoramidites 4Y and 4Z. DMTr = 4,4’-dimethoxytrityl; PCl reagent = 2-cyanoethyl-
N,N‘-diisopropylchlorophosphoramidite; 1-Py = pyren-1-yl; 2-Py = pyren-2-yl; 4-Py = pyren-4-yl.
nol23 to borane – afforded 2Y and 2Z in modest but acceptable
Monomers X–Z were studied in 9-mer sequence contexts,
yields (30% and 20% respectively, Scheme 1). Subsequent which we have previously used to screen and identify potential
O5′-dimethoxytritylation gave nucleosides 3Y and 3Z, which Invader building blocks.16 ONs containing a single incorpor-
upon treatment with 2-cyanoethyl-N,N-diisopropylchloropho- ation in the 5′-GBG ATA TGC context are denoted X1, Y1, and
sphoramidite (PCl reagent) and Hünig’s base provided target Z1. Similar conventions apply for the B2–B6 series (Table 1).
phosphoramidites 4Y and 4Z in excellent yield.
Reference DNA and RNA strands are denoted D1/D4 and
R1/R4, respectively (see footnote, Table 1).
Synthesis of modified ONs and experimental design
Thermostability of duplexes with complementary DNA/RNA
Phosphoramidites 4X, 4Y and 4Z were used in machine-
assisted solid-phase DNA synthesis to incorporate monomers First, the thermostabilities of duplexes between B1–B6 and
X–Z into ONs using 4,5-dicyanoimidazole as an activator and complementary DNA or RNA targets were determined from
extended hand-coupling (15 min), which resulted in stepwise thermal denaturation experiments performed in medium salt
coupling yields of ∼99%/∼99%/∼98% for 4X/4Y/4Z, respect- phosphate buffer ([Na+] = 110 mM, pH 7.0). The resulting
ively. The identity and purity of the modified ONs was estab- denaturation curves display the expected monophasic sigmoi-
lished via MALDI-TOF (Table S1 in the ESI†) and ion-pair dal transitions (Fig. S1 in the ESI†). Interestingly, Y-modified
reverse phase HPLC (>85% purity), respectively.
ONs form much more thermostable duplexes with cDNA than
Table 1 Thermal denaturation temperatures (Tm’s) for duplexes between B1–B6 and complementary DNA or RNAa
Tm (ΔTm/mod) [°C]
+cDNA
+cRNA
ON
Sequence
B =
Xb
Y
Z
Xb
Y
Z
B1
B2
B3
B4
B5
B6
5′-GB
5′-GTG AB
5′-GTG ATA B
3′-CAC BAT ACG
3′-CAC TAB
3′-CAC BAB
̲
34.5 [+5.0]
42.5 [+13.0]
37.5 [+8.0]
33.0 [+3.5]
42.5 [+13.0]
43.5 [+7.0]
35.5 [+6.0]
43.5 [+14.0]
39.0 [+9.5]
35.5 [+6.0]
43.5 [+14.0]
45.5 [+8.0]
29.5 [ 0.0]
37.5 [+8.0]
33.5 [+4.0]
26.5 [−3.0]
38.5 [+9.0]
32.5 [+1.5]
24.5 [−2.0]
30.5 [+4.0]
26.5 [ 0.0]
20.5 [−4.5]
27.5 [+2.5]
24.0 [−0.3]
25.5 [−1.5]
34.0 [+7.5]
28.0 [+1.5]
23.0 [−1.5]
32.0 [+7.5]
28.5 [+2.0]
20.5 [−6.0]
27.5 [+1.0]
21.5 [−5.0]
16.5 [−8.0]
26.5 [+2.0]
16.5 [−4.0]
̲
̲
̲
̲
ACG
ACG
̲
̲
a ΔTm = change in Tm relative to reference duplexes D1:D4 (Tm ≡ 29.5 °C), D1:R4 (Tm ≡ 26.5 °C) or R1:D4 (Tm ≡ 24.5 °C), where D1:
5′-GTGATATGC, D4: 3′-CACTATACG, R1: 5′-GUGAUAUGC and R4: 3′-CACUAUACG; Tm’s are determined as the maximum of the first derivative of
melting curves (A260 vs. T) recorded in medium salt buffer ([Na+] = 110 mM, [Cl−] = 100 mM, pH 7.0 (NaH2PO4/Na2HPO4)), using 1.0 µM of each
strand. Tm’s are averages of at least two measurements within 1.0 °C; A = adenin-9-yl DNA monomer, C = cytosin-1-yl DNA monomer, G = guanin-
9-yl DNA monomer, T = thymin-1-yl DNA monomer. For structures of monomers X–Z see Fig. 1. b From ref. 17b.
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studies in which reduced binding specificity of intercalator-
modified ONs was observed.26 Interestingly, the specificity
trends mirror the affinity trends. Thus, Y2 discriminates mis-
matched target more efficiently than X2, which, conversely,
discriminates mismatched targets more efficiently than Z2.
Additional specificity data are discussed in the ESI (Tables S3
and S4†).
Table 2 Discrimination of mismatched DNA targets by X2/Y2/Z2 and
reference strandsa
DNA: 3′-CAC TB
̲
Tm [°C]
ON
Sequence
B
̲
=
A
C
G
T
D1
X2b
Y2
5′-GTG ATA TGC
29.5
42.5
43.5
37.5
−16.5
−13.5
−16.5
−10.0
−9.5
−5.5
−6.5
−4.0
−17.0
−7.0
−9.0
−3.0
5′-GTG AX
5′-GTG AYA TGC
5′-GTG AZA TGC
̲A TGC
Optical spectroscopy
̲
Z2
̲
UV-Vis absorption and steady-state fluorescence emission
spectra of X/Y/Z-modified ONs in the presence or absence of
complementary DNA/RNA targets were recorded next to further
ascertain intercalative binding modes of the pyrene moieties,
as intercalation is known to induce bathochromic shifts of
pyrene absorption peaks27 and nucleobase-mediated quench-
ing of pyrene fluorescence.27a,28 Indeed, X/Y/Z-modified ONs
generally display hypochromic and bathochromic shifts in the
pyrene absorption spectra upon hybridization with DNA and
RNA targets (Δλmax = 0–5 nm, Table 3; Fig. S2–S4 in the ESI†).
Slightly greater bathochromic shifts are generally observed
upon hybridization with DNA than RNA targets, which again
may reflect the preference of intercalators for the less com-
pressed geometry of B-DNA.
Steady-state fluorescence emission spectra of X/Y/Z-modi-
fied ONs and the corresponding duplexes with cDNA/cRNA
display two vibronic bands at λem = 382 3 nm and 402
3 nm, respectively, as well as a small shoulder at ∼420 nm. As
anticipated, the fluorescence intensity typically decreases
upon hybridization with DNA/RNA targets, with greater
decreases typically being observed upon DNA binding (Fig. 2;
Fig. S5–S7 in the ESI†). Interestingly, duplexes modified with
monomer Z are noticeably less fluorescent than X- or Y-modi-
fied duplexes.
a For conditions of thermal denaturation experiments, see Table 1.
Tm’s of fully matched duplexes are shown in bold. ΔTm = change in Tm
relative to fully matched duplex. b From ref. 17b.
unmodified ONs (ΔTm/mod = +6.0 to +14.0 °C, Table 1). In
fact, the resulting duplexes are even more thermostable than
X-modified duplexes, suggesting that the 2-pyrenyl moiety of
monomer Y is very well accommodated in DNA duplexes. In
contrast, Z-modified ONs display considerably lower affinity
toward cDNA (ΔTm/mod = −3.0 to +9.0 °C, Table 1). The
thermostability trends are sequence-dependent. Thus, ONs in
which the pyrene-functionalized monomers are flanked by
3′-purines, induce greater stabilization than ONs with 3′-flank-
ing pyrimidines (e.g., compare ΔTm/mod values for B2- and
B4-series, Table 1). This is indicative of 3′-intercalative binding
modes of the pyrene moieties,19,24 as 3′-flanking purines
would provide larger π-stacking surfaces than 3′-pyrimidines.
Less stable duplexes are formed with cRNA (ΔTm/mod = −8.0
to +7.5 °C, Table 1; trend: Y > X > Z), which also points to inter-
calative pyrene binding modes,18,24a,25 as intercalators gener-
ally favor the less compressed B-type helix geometry of DNA:
DNA duplexes.26
Binding specificity
Molecular modeling of probe–target duplexes
Next, the binding specificity of singly modified ONs (B2-series)
was studied using DNA targets with mismatched nucleotides To rationalize the observed biophysical trends, we performed
opposite of the pyrene-functionalized monomer (Table 2). force-field calculations on duplexes between X2/Y2/Z2 and
X2/Y2/Z2 discriminate mismatched DNA targets less efficiently complementary DNA (residue numbering: 5′-G1T2G3A4B5A6-
than reference strand D1, especially when a mismatched T is T7G8C9:3′-C18A17C16T15A14T13A12C11G10). The complete data set
opposite of the modification. This is in agreement with other of structural furanose, base pair and dinucleotide step para-
Table 3 Absorption maxima in the 335–355 nm region for single-stranded X/Y/Z-modified ONs and the corresponding duplexes with complemen-
tary DNA or RNAa
λmax [Δλmax]/nm
Xb
Y
Z
ON
Sequence
B̲
=
SSP
+cDNA
+cRNA
SSP
+cDNA
+cRNA
SSP
+cDNA
+cRNA
B1
B2
B3
B4
B5
5′-GB
5′-GTG AB
5′-GTG ATA B
3′-CAC BAT ACG
3′-CAC TAB ACG
̲
350
348
350
350
349
353[+3]
353[+5]
353[+3]
352[+2]
353[+4]
352[+2]
352[+4]
352[+2]
352[+2]
352[+3]
344
342
344
344
343
346[+2]
345[+3]
346[+2]
345[+1]
345[+2]
345[+1]
345[+3]
345[+1]
345[+1]
345[+2]
343
340
343
342
342
344[+1]
343[+3]
344[+1]
344[+2]
344[+2]
343[ 0]
343[+3]
343[ 0]
343[+1]
344[+2]
̲
̲
̲
̲
a Measurements were performed at 5 °C using a spectrophotometer and quartz optical cells with 1.0 cm path lengths. Buffer conditions are as for
thermal denaturation experiments. b From ref. 17b.
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Fig. 2 Steady-state fluorescence emission spectra of select X/Y/Z-modified ONs and corresponding duplexes with DNA/RNA targets. Spectra were
recorded at T = 5 °C using λex = 350, 345 and 340 nm for X, Y and Z-modified ONs, respectively. Each strand was used at 1.0 μM concentration in Tm
buffer.
meters29 is provided in the ESI (Tables S7–S8 and Fig. S9– tates stacking with all four neighboring nucleobase moieties
S14†).
(Fig. 3 bottom). Conversely, the pyrene of monomer Z stacks
In agreement with the photophysical data presented above, only with the flanking nucleobases on its own strand, i.e., Z5
the pyrene moieties remain stably intercalated in the duplex and A6. The poorer fit of the pyrene of monomer Z is also
core throughout the stochastic dynamics simulations (Fig. 3 reflected in greater perturbation across the B5A6:A14T13 di-
top – for details of calculation protocol, see Experimental nucleotide step, i.e., more pronounced buckle (∼6° and ∼−4° in
section). Inspection of the structures reveals that the right- D1:D4, ∼13° and ∼−10° in X2:D4, ∼15° and ∼−11° in Y2:D4,
handed duplexes are elongated relative to the corresponding and ∼21° and ∼−19° in Z2:D4, Fig. S9 in the ESI†) and
B-DNA reference duplex (e.g., rise increases from ∼3.3 Å to decreased roll (∼12° in D1:D4, ∼−10° in X2:D4, ∼−11° in Y2:
∼7.3 Å across the B5A6:A14T13 dinucleotide step, Fig. S10 in the D4, and ∼−19° in Z2:D4, Fig. S10 in the ESI†). Another note-
ESI†). The overlap between the pyrene and neighboring worthy observation is that the furanose rings of nucleotides
nucleobases is strongly influenced by the substitution pattern X5A6, Y5A6 and Z5A6 generally exhibit increased South type
of the pyrene, which explains the observed thermostability character relative to the reference duplex (P = 124–171° vs.
trends (Tm decreases Y ≥ X > Z). The pyrene of monomer X 102–129°), presumably as a consequence of π–π-stacking
stacks with the nucleobase moieties of X5, A6 and A13, while between the pyrene and flanking nucleobases (Table S7 in the
the pyrene of monomer Y is positioned in a manner that facili- ESI†).
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Fig. 3 Lowest energy structures of X2:D4 (left), Y2:D4 (middle) and Z2:D4 (right). Upper: side view of duplex; lower: top view of the central duplex
region. Color code: sugar phosphate backbone (red); pyrene moieties (blue) and nucleobases (green). Hydrogen atoms, sodium ions and bond
orders are omitted for clarity.
To sum up, the molecular modeling structures are useful in probe duplexes, Table 4). Clearly the two pyrene-functionalized
explaining the observed thermostability trends and photo- monomers interact with each other in an energetically unfavor-
physical properties of X/Y/Z-modified duplexes.
able manner when placed in this motif. Interestingly, there is
little difference in the dsDNA-targeting potential of X2:X5,
Y2:Y5 and Z2:Z5 as judged by the DA values; the lower thermo-
stability of Z-modified probe–target duplexes is compensated
by an equivalent destabilization of the probe duplexes
(compare DA values for X2:X5, Y2:Y5 and Z2:Z5, Table 4).
The above Tm-based conclusions were corroborated through
analysis of thermodynamic parameters for duplex formation,
which were derived through fitting of thermal denaturation
curves.30 Thus, formation of probe–target duplexes is highly
favorable relative to unmodified duplexes, with Y-modified
and Z-modified ONs leading to the most and least stable
duplexes with cDNA, respectively (see the first two ΔΔG293
columns, Table 4). Stabilization of the probe–target duplexes is
largely a consequence of increased entropy (Tables S5 and S6
in the ESI†). On the other hand, B2:B5 duplexes are much less
stable (see the third ΔΔG293 column, Table 4) due to strongly
increased enthalpy (Tables S5 and S6 in the ESI†). Consequen-
Biophysical properties of duplexes with interstrand zippers of
X/Y/Z-monomers
Next, we determined the thermostability of DNA duplexes with
different interstrand zipper arrangements of X/Y/Z monomers
to identify monomers and probe architectures that show
the greatest potential for dsDNA-recognition. The impact on
duplex thermostability upon incorporation of
a second
monomer can be additive, greater-than-additive or less-than-
additive relative to the corresponding singly modified
duplex. The impact is conveniently approximated in terms
of Tm’s by the term ‘deviation from additivity’ (DA) defined as:
DAONA:ONB ≡ ΔTm (ONA:ONB) − [ΔTm (ONA:cDNA) + ΔTm
(cDNA:ONB)], where ONA:ONB is a duplex with an interstrand
arrangement of monomers. DA also serves as an indicator for
dsDNA-recognition potential. Probes with strongly negative
DA values are likely to be activated for recognition of iso-
sequential dsDNA via the process depicted in Fig. 1, since the
products of the recognition process (i.e., probe–target
duplexes) are more thermostable than the reactants (i.e., probe
duplexes and target duplexes). A more rigorous approach
based on differences in ΔG values of probe–target and Invader
duplexes is discussed shortly.
tially, B2:B5 probes display favorable binding free energy for
293
recognition of iso-sequential dsDNA targets as given by ΔG
rec
(ONA:ONB) = ΔG293 (ONA:cDNA) + ΔG293 (cDNA:ONB) − ΔG293
(ONA:ONB) − ΔG293 (dsDNA) (i.e., ΔG
for B2:B5 ≪ 0 kJ
293
rec
293
mol−1, Table 4). X2:X5 and Y2:Y5 display very similar ΔG
rec
values. Unfortunately, the absence of a clear lower base line in
As expected,13,15,16 duplexes with +1 interstrand monomer
arrangements are less thermostable and more strongly acti-
vated for dsDNA-recognition than duplexes with other arrange-
ments (compare Tm’s and DA values for B2:B5 relative to other
the thermal denaturation curve of Z2:Z5 precluded determi-
293
nation of ΔG
for this duplex. Probes with other interstrand
rec
arrangements of X/Y/Z-monomers are much more stable and
display much less favorable binding free energy for recognition
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Table 4 Properties of X/Y/Z-modified probe duplexesa
ΔG293 [ΔΔG293] (kJ mol−1
)
ON ZP
Sequence
Tm (°C) DA (°C) Upper ON vs. cDNA Lower ON vs. cDNA Probe duplex
ΔG2re9c3 (kJ mol−1
)
λmax (nm)
X1
+4
X5
5′-GX
3′-CAC TAX
5′-GXG ATA TGC
3′-CAC XAT ACG
5′-GTG AXA TGC
3′-CAC TAX ACG
5′-GTG AXA TGC
3′-CAC XAT ACG
5′-GTG ATA XGC
3′-CAC TAX ACG
5′-GTG ATA XGC
3′-CAC XAT ACG
̲
47.5
31.5
26.5
39.5
45.5
40.5
0.0
−6.5
−29.0
−6.5
−5.0
−0.5
−46 0 [−5]
−46 0 [−5]
−55 1 [−14]
−54 0 [−13]
−50 0 [−9]
−50 0 [−7]
−52 0 [−11]
−46 0 [−5]
−53 1 [−12]
−46 0 [−5]
−52 0 [−11]
−46 0 [−3]
−55 1 [−14]
−45 1 [−4]
−38 1 [+3]
−51 1 [−10]
−55 1 [−14]
−53 1 [−10]
−2
−6
−29
−8
−6
0
352
350
348
351
352
352
̲
X1
+2
X4
̲
̲
X2
+1
X5
̲
̲
X2
−1
X4
̲
̲
X3
−1
X5
̲
̲
X3
−3
X4
̲
̲
Y1
+4
Y5
5′-GY
3′-CAC TAY
5′-GYG ATA TGC
3′-CAC YAT ACG
5′-GTG AYA TGC
3′-CAC TAY ACG
5′-GTG AYA TGC
3′-CAC YAT ACG
5′-GTG ATA YGC
3′-CAC TAY ACG
5′-GTG ATA YGC
3′-CAC YAT ACG
̲
48.5
35.5
28.5
43.5
47.5
43.0
−1.0
−6.0
−29.5
−6.0
−5.5
−2.0
−48 0 [−7]
−48 0 [−7]
−56 0 [−15]
−56 0 [−15]
−51 1 [−10]
−51 1 [−10]
−55 1 [−14]
−46 0 [−5]
−55 1 [−14]
−46 0 [−5]
−55 1 [−14]
−46 0 [−5]
−56 1 [−15]
−45 1 [−4]
−43 1 [−2]
−55 1 [−14]
−58 1 [−17]
−53 1 [−12]
−6
−8
346
346
339
346
346
346
̲
Y1
+2
Y4
̲
̲
Y2
+1
Y5
̲
−27
−6
̲
Y2
−1
Y4
̲
̲
Y3
−1
Y5
̲
−7
̲
Y3
−3
Y4
̲
−3
̲
Z1
+4
Z5
5′-GZ
3′-CAC TAZ
5′-GZG ATA TGC
3′-CAC ZAT ACG
5′-GTG AZA TGC
3′-CAC TAZ ACG
5′-GTG AZA TGC
3′-CAC ZAT ACG
5′-GTG ATA ZGC
3′-CAC TAZ ACG
5′-GTG ATA ZGC
3′-CAC ZAT ACG
̲
38.5
27.5
19.5
30.5
37.0
32.5
0.0
+1.0
−44 0 [−3]
−44 0 [−3]
−48 1 [−7]
−48 1 [−7]
−46 0 [−5]
−46 0 [−5]
−48 1 [−7]
−41 0 [ 0]
−48 1 [−7]
−41 0 [ 0]
−48 1 [−7]
−41 0 [ 0]
−49 1 [−8]
−41 1 [ 0]
N/A
−2
−3
345
342
339
344
345
345
̲
Z1
+2
Z4
̲
̲
Z2
+1
Z5
̲
−27.0
−4.0
−5.5
+2.0
N/A
−7
̲
Z2
−1
Z4
̲
−41 1 [ 0]
−46 1 [−5]
−42 1 [−1]
̲
Z3
−1
Z5
̲
−7
̲
Z3
−3
Z4
̲
−4
̲
a ZP = zipper. For conditions of thermal denaturation and absorption experiments, see Tables 1 and 3, respectively. DA = ΔTm (Invader) − (ΔTm
293
(upper strand vs. cDNA) + ΔTm (lower strand vs. cDNA)). ΔΔG293 is measured relative to ΔG293 for D1:D4 = −41 kJ mol−1. ΔG = ΔG293 (upper
rec
strand vs. cDNA) + ΔG293 (lower strand vs. cDNA) − ΔG293 (probe duplex) − ΔG293 (dsDNA target). “ ” denotes standard deviation. N/A = the
absence of a clear lower base line prevented determination of this value. Tm’s for X-modified duplexes have been previously published in ref. 17b
but are included to facilitate direct comparison.
293
of iso-sequential dsDNA targets (ΔG
between−8 and 0 kJ and unstructured emission centered at λem ∼ 490 nm, which is
rec
mol−1, Table 4), presumably since the two monomers act inde- consistent with pyrene excimer emission (Fig. 4).15,16,19,31 The
pendently from each other. presence of these signals shows that the pyrene moieties are
The unique characteristics of duplexes with +1 interstrand stacking with an interplanar separation of ∼3.4 Å. We specu-
monomer arrangements relative to other probe duplexes are also late that stacking occurs in the major groove in a similar
reflected in the blue-shifted pyrene absorption peaks (Fig. S8 in manner as previously suggested for duplexes with +2 zipper
the ESI†), which are indicative of reduced pyrene–nucleobase arrangements of pyrene-functionalized ara-uridine mono-
interactions (compare λmax for B2:B5 with λmax for other probe mers.31b,32 Duplexes with +1 interstrand monomer arrange-
duplexes (Table 4) or probe–target duplexes (Table 3)).
ments also display excimer emission albeit of much weaker
intensity. In this case, the two pyrene moieties most likely
All probe duplexes exhibit vibronic peaks at λem = 380
2 nm, 400
3 nm and ∼420
3 nm in their steady-state stack inside the duplex core in a similar manner as recently
fluorescence emission spectrum (Fig. 4). In addition, duplexes suggested for duplexes modified with 2′-N-(pyren-1-yl)methyl-
with +2 interstrand monomer arrangements display prominent 2′-amino-α-L-LNA monomers; the spatial separation for
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Fig. 4 Steady-state fluorescence emission spectra of duplexes with different interstrand monomer arrangements of X, Y, and Z (zipper type indi-
cated in parenthesis). For experimental conditions, see Fig. 2. Spectra for X-modified duplexes were previously reported in ref. 16 but are included
for comparison.
pyrene–pyrene stacking in the major groove is too large.15a,16 duplexes; base pairing in the vicinity of the interstrand
Dual intercalation would also explain the energetic lability of zipper is distorted in both structures and the presence of a
the B2:B5 duplexes, as a localized region with one intercalator large hydrophobic pyrene in the major grove (Type B) will
per base pair would ensue, which represents a violation of the almost certainly perturb the stabilizing hydration layer or
‘nearest neighbor exclusion principle’.14
cause unfavorable steric interactions.
Molecular modeling of B2:B5 duplexes
Recognition of DNA hairpins using activated probe duplexes
293
rec
Force-field based simulations of B2:B5 duplexes were The ΔG
values indicate that duplexes with +1 interstrand
performed to gain additional insight into the binding mode zippers of X/Y/Z monomers are the most strongly activated
of the pyrene moieties. The results of these simulations probes for dsDNA-recognition (Table 4). We therefore decided
point toward two possible binding modes, i.e., Type A in which to evaluate these probes in greater detail using a gel shift assay
both pyrene moieties intercalate into the duplex core (Fig. 5) that we have used to screen other potential Invader monomers
and Type B in which one pyrene is intercalating and the other with.13,16 Thus, a digoxigenin (DIG) labeled DNA hairpin (DH)
is extruded into the major groove (Fig. S15 in the ESI†). The – comprised of a 9-mer double-stranded mixed-sequence stem
presence of excimer signals in the steady state emission that is linked by a T10 loop – was used as a model target
spectra of X2:X5 and Z2:Z5 (Fig. 4), suggests that these (Fig. 6a).33 As expected, the unimolecular nature of DH1
duplexes are capable of adopting Type A conformations. strongly stabilizes the stem region as seen from the signifi-
However, the weak intensity of the excimer signals – especially cantly higher Tm relative to the corresponding linear DNA
with Y2:Y5 – suggests that B2:B5 duplexes equilibrate between duplex (Tm = 57.0 °C vs. 29.5 °C, respectively; Fig. 6b and
Type A and Type B conformations. Type A structures are Table 1). Incubation of DH1 with X2:X5 or Y2:Y5 in a HEPES
characterized by significant elongation (rise ∼ 10 Å) and local buffer for 15 h at ambient temperature resulted in dose-depen-
perturbation (buckle ∼ 30° and ∼−30° at B5A6 and A14B13, dent recognition as evidenced by the emergence of a slower
respectively, and roll ∼ −30° across X5A6:X14B13 and Y5A6: migrating band on non-denaturing PAGE gels (Fig. 6c).33 It is
Y14B13 and ∼−18° across Z5A6:A14) (Fig. S11–S12 in the ESI†). particularly noteworthy that as little as 1.0 molar equivalent of
Type B structures are less elongated (rise ∼ 7.5 Å) and less per- Y2:Y5 resulted in ∼50% dsDNA-recognition (Fig. 6d). In con-
turbed (buckle ∼ 20° and ∼−20° at B5A6 and A14B13, respect- trast, very little recognition was observed even at 500-fold
ively, and roll ∼ −5°) (Fig. S13–S14 in the ESI†). Both molar excess of Z2:Z5 (Fig. 6c and 6d). This was surprising to
conformations can account for the lability of the B2:B5 us considering the very similar DA values of the B2:B5
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Fig. 5 Lowest energy structures of X2:X5 (left), Y2:Y5 (middle) and Z2:Z5 (right) in Type A conformation. Top: side view of duplex; bottom: top view
of the central duplex region. For color code, see Fig. 3.
duplexes (Table 4). To determine if the poor dsDNA-recog-
nition efficiency of Z2:Z5 was due to insufficient stability of
Conclusion
the recognition complex and/or slow reaction kinetics, we Short synthetic routes to suitably protected 2′-O-(pyren-2-yl)-
performed separate experiments in which DH1 was annealed methyluridine and 2′-O-(pyren-4-yl)methyluridine have been
in the presence of 500-fold molar excess of X2:X5 or Z2:Z5 developed. ONs modified with 2′-O-(pyren-2-yl)methyluridines
(i.e., heated to 95 °C for 2 min, cooled to 8° over 3 h, then ana- display greater affinity toward complementary DNA and better mis-
lyzed on non-denaturing PAGE gels). The resulting gel electro- match discrimination than ONs modified with the corresponding
phoretograms (results not shown) were essentially identical to 1-pyrenyl or 4-pyrenyl analogues. Molecular modeling suggests
those from the room temperature incubation experiments dis- this to be a consequence of more efficient π–π-stacking between
cussed above, which suggests that the recognition complex the intercalating pyrene moiety and neighboring base pairs.
between Z2:Z5 and DH1 is not sufficiently stable at the experi-
mental conditions of the assay.
DNA duplexes with +1 interstrand arrangements of these
pyrene-functionalized monomers display lower thermostabil-
As expected, the unmodified control duplex D1:D4 did not ity, more blue-shifted pyrene absorption maxima, more dis-
result in formation of recognition complexes even when used tinctive fluorescence emission profiles and greater potential
at 500-fold molar excess, as there is no driving force to over- for dsDNA-recognition than duplexes with other monomer
come the energetic penalty of opening DH1 (Fig. 6c). The use arrangements. Force field simulations suggest that the +1
of 500-fold molar excess of single-stranded X2/X5/Y2/Y5 only zipper motif results in significant perturbation of the duplex.
resulted in 15–40% recognition of DH1 (Fig. 6e), which Although the chemical differences between the three studied
demonstrates that both probe strands are necessary for monomers are relatively minor, significant variations in the
efficient dsDNA-recognition.
dsDNA-recognition efficiencies of the resulting probes are
Finally, the binding specificity of X2:X5 and Y2:Y5 was observed. Thus, probes with +1 interstrand zippers of 2′-O-
examined in detail by incubating the probe duplexes with (pyren-2-yl)methyluridines recognize mixed-sequence DNA
DNA hairpins DH2–DH7, which are fully base-paired but hairpins very efficiently, whereas probes constructed with
singly mismatched relative to the Invader probes (underlined the corresponding 2′-O-(pyren-4-yl)methyluridines do not.
residues indicate position of sequence deviation, Fig. 6b). Remarkably, Invaders based on 2′-O-(pyren-2-yl)methyluri-
Importantly, neither a 500-fold molar excess of X2:X5 nor Y2: dines result in ∼50% recognition of dsDNA when used in equi-
Y5 resulted in recognition of the singly mismatched DNA hair- molar quantities. These findings encourage additional
pins, which demonstrates that hairpin recognition is highly structure–property relationship studies based on the 2-pyrenyl
specific (Fig. 6f).
scaffolds with the goal of further increasing the dsDNA recog-
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Fig. 6 Recognition of model dsDNA targets using activated probe duplexes. (a) Illustration of recognition process; (b) sequences and intramolecular
m’s of DNA hairpins with isosequential (DH1) or mismatched stems (DH2-DH7) relative to B2:B5 probes (for conditions of thermal denaturation
T
experiments, see Table 1); (c) representative gel electrophoretograms illustrating recognition of DH1 using 1- to 500-fold molar excess of X2:X5,
Y2:Y5, Z2:Z5, or unmodified D1:D4; (d) dose-response curves (average of three independent experiments; error bars represent standard deviation);
(e) incubation of DH1 with 500-fold molar excess of single-stranded X2, X5, Y2 or Y5 or double-stranded X2:X5 or Y2:Y5; (f) gel electrophoreto-
grams illustrating incubation of DH1–DH7 with 500-fold molar excess of X2:X5 or Y2:Y5. Experimental conditions for electrophoretic mobility shift
assay: separately pre-annealed targets (34.4 nM) and probes (variable concentration) were incubated for 15 h at ambient temperature in 1× HEPES
buffer (50 mM HEPES, 100 mM NaCl, 5 mM MgCl2, 10% sucrose, 1.4 mM spermine tetrahydrochloride, pH 7.2) and then run on 16% non-denaturing
PAGE (performed at 70 V, 2 h, ∼4 °C) using 0.5× TBE as a running buffer (45 mM Tris, 45 mM boric acid, 1 mM EDTA); DIG: digoxigenin.
nition efficiency of Invader probes, especially considering that (petroleum ether) to provide an off-white solid material
detection of chromosomal DNA using Invaders based on the (5.50 g), which was used in the next step without further
slightly less efficient 2′-O-(pyren-1-yl)-RNA monomers already purification.
has been demonstrated.13 Proof-of-concept studies aiming at
The solid material was dissolved in anhydrous benzene
expanding the repertoire of applications for Invader probes in (400 mL) along with DDQ (14.4 g, 63.6 mmol), and the solu-
molecular biology, nucleic acid diagnostics and medicinal tion was refluxed under an argon atmosphere for 4 h. The
chemistry are ongoing and will be reported in due course.
resulting slurry was filtered through a pad of celite, which was
thoroughly washed with benzene. The filtrate was washed with
aq. NaOH (10% v/v, 3 × 100 mL) and water (3 × 100 mL). The
organic phase was evaporated to dryness and the resulting
residue purified by silica gel column chromatography (pet-
roleum ether) to afford a pale yellow solid material (4.60 g),
which was used in the next step without further purification.
Next, n-butyllithium (9.82 mL, 24.5 mmol, 2.5 M in hexane)
was added slowly to a −78 °C solution of the solid material in
anhydrous ether–THF (350 mL, 1 : 1 v/v). The brightly red solu-
tion was stirred at −78 °C for 2 h and then slowly treated with
anhydrous DMF (2.6 mL, 33.5 mmol). The reaction mixture
was stirred at −30 to −50 °C for 1 h and then at rt for an
additional 15 h. At this point, the reaction mixture was poured
into ice-cold water (∼100 mL) and the aqueous phase was
Experimental
Preparation of 2-pyrenemethanol
2-Pyrenemethanol was obtained from 4,5,9,10-tetrahydro-
pyrene by combining known literature protocols.22 First, a
solution of bromine (1.24 mL, 24.2 mmol) in anhydrous DMF
(40 mL) was added dropwise over 2 h to a room temperature
solution of 4,5,9,10-tetrahydropyrene22a (5.00 g, 24.2 mmol) in
anhydrous DMF (40 mL). The mixture was stirred for 4 h at rt
after completed addition, poured into cold water and stirred
overnight. The resulting precipitate was filtered, washed with
water, dried and purified by silica gel column chromatography
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extracted with EtOAc (3 × 100 mL). The combined organic column chromatography (0–3% CH2Cl2 in pet ether, v/v) to
layers were evaporated to near dryness and the resulting afford a brightly yellow solid material (1.20 g), which was used
residue was purified by silica gel column chromatography in the next step without further purification.
(0–3% CH2Cl2 in petroleum ether, v/v) to afford a brightly
yellow solid material (3.50 g), which was used in the next step of the solid material in anhydrous THF (80 mL) and the reac-
without further purification. tion mixture was stirred overnight at rt. The reaction mixture
Finally, NaBH4 (0.24 g, 6.25 mmol) was added to a solution
Lastly, sodium borohydride (0.69 g, 18.2 mmol) was added was cooled on an ice-bath and aqueous NaHCO3 (10% v/v,
to a solution of this solid material in anhydrous THF (120 mL) 20 mL) was added carefully. The aqueous layer was extracted
and the reaction mixture was stirred overnight at rt. The reac- with EtOAc (3 × 80 mL) and the combined organic layers were
tion mixture was cooled on an ice bath and aqueous NaHCO3 evaporated to dryness. The resulting residue was purified by
(10% v/v, 50 mL) was added carefully. The aqueous layer was silica gel column chromatography (0–1% MeOH in CH2Cl2,
extracted with EtOAc (3 × 100 mL) and the combined organic v/v) to afford 4-pyrenemethanol (1.15 g, 41% over four steps) as
layers were evaporated to dryness. The resulting residue was a white solid material. Rf = 0.5 (10% MeOH in CH2Cl2, v/v);
purified by silica gel column chromatography (0–1% MeOH in MALDI-HRMS m/z 232.0881 ([M]+, C17H12O, Calc. 232.0888);
CH2Cl2, v/v) to afford 2-pyrenemethanol22c (3.45 g, 62% over 1H NMR (DMSO-d6): δ 8.39–8.41 (dd, 1H, J = 7.4 Hz, 0.8 Hz,
four steps) as a white solid material. Rf = 0.5 (10% MeOH in H3), 8.26–8.32 (m, 3H, H1/H6/H8), 8.24 (s, 1H, H5), 8.17–8.21
CH2Cl2, v/v); MALDI-HRMS m/z 232.0875 ([M]+, C17H12O, Calc. (2d, 2H, J = 8.8 Hz, H9/H10), 8.05–8.11 (m, 2H, H2/H7), 5.53 (t,
232.0888); 1H NMR (DMSO-d6): δ 8.28 (d, 2H, J = 7.8 Hz, 1H, ex, J = 5.5 Hz, OH), 5.19–5.21 (dd, 2H, J = 5.5 Hz, 0.8 Hz,
H6/H8), 8.25 (s, 2H, H1/H3), 8.17 (s, 4H, H4/H5/H9/H10), CH2Py); 13C NMR (DMSO-d6) δ 137.1, 130.9, 130.4, 130.3,
8.03–8.06 (ap t, 1H, J = 7.5 Hz, H7), 5.52 (t, 1H, ex, J = 5.7 Hz, 128.9, 127.4 (C9/C10), 127.1 (C9/C10), 126.2 (C2/C7), 125.9
OH), 4.97 (d, 2H, J = 5.7 Hz, CH2OH); 13C NMR (DMSO-d6) (C2/C7), 125.1 (C1/C6/C8), 125.0 (C1/C6/C8), 124.8 (C1/C6/C8),
δ 140.7 (C2), 130.5, 130.4, 127.31 (C4/C5),34 127.29 (C9/C10), 124.3 (C5), 124.0, 123.2, 121.2 (C3), 61.4 (CH2Py).
125.9 (C7), 124.9 (C6/C8), 123.8, 123.0 (C1/C3), 122.9, 63.2
General O2′-alkylation protocol for the preparation of 2Y/2Z
(description for ∼4.4 mmol scale)
(CH2Py).
Preparation of 4-pyrenemethanol
The appropriate pyrenemethanol, solid NaHCO3 and borane
4-Pyrenemethanol was obtained from 4,5,9,10-tetrahydro- (1.0 M solution in THF) were placed in a pressure tube, sus-
pyrene by combining known literature protocols23 and protocols pended in anhydrous DMSO and stirred under an argon
for the synthesis of 2-pyrenemethanol. First, a solution of atmosphere at rt until effervescence ceased (∼10 min). At this
bromine (0.62 mL, 12.0 mmol) in glacial acetic acid (25 mL) point, O2,O2′-anhydrouridine 120 was added (specific quan-
was added dropwise over 1 h to a room temperature solution tities of substrates and reagents are given below), the pressure
of 4,5,9,10-tetrahydropyrene 322a (2.50 g, 12.0 mmol) in glacial tube was purged with argon and sealed, and the mixture was
acetic acid (25 mL). After ended addition, the reaction mixture heated at ∼140 °C until analytical TLC indicated full conver-
was heated to 80 °C for 30 min, and then slowly cooled to rt, sion (∼3 h). After cooling to rt, the reaction mixture was
leading to the formation of a precipitate, which was isolated to poured into water (∼50 mL), stirred for 30 min and diluted
provide a white solid material (2.90 g, 10.1 mmol) which used with EtOAc (∼100 mL). The organic phase was washed with
in the next step without further purification.
water (4 × 50 mL), evaporated to dryness and the resulting
The solid material and DDQ (7.56 g, 33.3 mmol) were dis- crude purified by silica gel column chromatography (2–4%,
solved in anhydrous benzene (300 mL) and refluxed for 4 h. MeOH in CH2Cl2, v/v) to afford a residue, which was further
The resulting slurry was filtered through a celite pad, which purified through precipitation from refluxing methanol
was thoroughly washed with benzene. The filtrate was washed (30 min) to afford nucleoside 2 (yield specified below).
with aq. NaOH (10% v/v, 3 × 80 mL) and water (3 × 80 mL).
2′-O-(Pyren-2-yl)methyl-uridine (2Y)
The organic phase was evaporated to dryness and the resulting
residue purified via silica gel column chromatography (pet- O2,O2′-Anhydrouridine 1 (1.00 g, 4.42 mmol), 2-pyrenemetha-
roleum ether) to obtain a pale yellow solid (2.00 g), which was nol (2.05 g, 8.84 mmol), NaHCO3 (74.3 mg. 0.88 mmol), BH3
used in the next step without further purification.
in THF (2.21 mL, 2.21 mmol) and anhydrous DMSO (8 mL)
Next, n-butyllithium (4.27 mL, 10.7 mmol, 2.5 M in hexane) were mixed, reacted, worked up, and purified as described
was added slowly to a −78 °C solution of this solid material in above to afford nucleoside 2Y (0.60 g, 30%) as a white solid
anhydrous ether–THF (170 mL, 1 : 1 v/v). The brightly red solu- material. Rf: 0.4 (10% MeOH in CH2Cl2, v/v); MALDI-HRMS
tion was stirred at −78 °C for 2 h and then treated slowly with m/z 481.1386 ([M + Na]+, C26H22N2O6·Na+, Calc. 481.1370;
anhydrous DMF (1.12 mL, 14.6 mmol). The reaction mixture 1H NMR (DMSO-d6):35 δ 11.38 (s, 1H, ex, NH), 8.28–8.30 (d, 2H,
was stirred at −30 to −50 °C for 1 h and then at rt for J = 7.8 Hz, H6Py, H8Py), 8.23 (s, 2H, H1Py, H3Py), 8.17–8.20 (d,
additional 15 h. The reaction mixture was poured into ice-cold 2H, J = 8.8 Hz, H4Py, H10Py), 8.10–8.13 (d, 2H, J = 8.8 Hz, H5Py,
water (∼50 mL), the aqueous phase was extracted with EtOAc H9Py), 8.07 (t, 1H, J = 7.8 Hz, H7Py), 7.85 (d, 1H, J = 8.0 Hz, H6),
(3 × 50 mL), and the combined organic phase evaporated to 6.09 (d, 1H, J = 4.9 Hz, H1′), 5.51 (d, 1H, J = 8.0 Hz, H5), 5.35
dryness. The resulting residue was purified by silica gel (d, 1H, ex, J = 5.7 Hz, 3′-OH), 5.16–5.19 (d, 1H, J = 12.7 Hz,
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CH2Py), 5.11 (t, 1H, ex, J = 4.9 Hz, 5′-OH), 5.03–5.06 (d, 1H, J = H3Py, H6Py, H8Py), 8.17–8.20 (d, 2H, J = 8.9 Hz, H4Py, H10Py),
12.7 Hz, CH2Py), 4.23–4.26 (m, 1H, H3′), 4.12–4.15 (m, 1H, 8.11–8.13 (d, 2H, J = 8.9 Hz, H5Py, H9Py), 8.07 (t, 1H, J = 7.5 Hz,
H2′), 3.98–4.00 (m, 1H, H4′), 3.65–3.69 (m, 1H, H5′), 3.57–3.62 H7Py), 7.67 (d, 1H, J = 8.0 Hz, H6), 7.31–7.33 (m, 2H, DMTr),
(m, 1H, H5′); 13C NMR (DMSO-d6): δ 163.0, 150.6, 140.2 (C6), 7.22–7.26 (m, 2H, DMTr), 7.15–7.20 (m, 5H, DMTr), 6.78–6.84
136.3, 130.54, 130.50, 127.5 (C4Py, C10Py), 127.2 (C5Py, C9Py), (m, 4H, DMTr), 6.07–6.08 (d, 1H, J = 3.9 Hz, H1′), 5.45 (d, 1H,
126.1 (C7Py), 125.1 (C6Py, C8Py), 123.7 (C1Py, C3Py), 123.2, 101.8 ex, J = 6.7 Hz, 3′-OH), 5.13–5.23 (m, 3H, H5, CH2Py), 4.31–4.35
(C5), 86.4 (C1′), 85.2 (C4′), 80.8 (C2′), 71.3 (CH2Py), 68.4 (C3′), (m, 1H, H3′), 4.18–4.20 (m, 1H, H2′), 4.09–4.12 (m, 1H, H4′),
60.5 (C5′).
3.68 (s, 3H, CH3O), 3.65 (s, 3H, CH3O), 3.30–3.34 (m, 1H, H5′;
overlap with H2O), 3.23–3.26 (m, 1H, H5′); 13C NMR
(DMSO-d6): δ 162.9, 158.1, 158.0, 150.4, 144.4, 140.1 (C6),
2′-O-(Pyren-4-yl)methyl-uridine (2Z)
O2,O2′-Anhydrouridine 1 (0.50 g, 2.21 mmol), 4-pyrenemetha- 136.2, 135.3, 135.0, 130.6, 130.5, 129.7 (DMTr), 129.6 (DMTr),
nol (1.03 g, 4.42 mmol), NaHCO3 (37.1 mg. 0.44 mmol), BH3 127.8 (DMTr), 127.6 (DMTr), 127.5 (Py), 127.2 (Py), 126.7 (Py),
in THF (1.10 mL, 1.10 mmol) and anhydrous DMSO (5 mL) 126.1 (Py), 125.1 (Py), 123.68, 123.66 (Py), 123.2, 113.2 (DMTr),
were mixed, reacted, worked up, and purified as described 113.1 (DMTr), 101.5 (C5), 87.2 (C1′), 85.9, 82.9 (C4′), 80.6 (C2′),
above to afford nucleoside 2Y (200 mg, 20%) as a white solid. 71.3 (CH2Py), 68.7 (C3′), 62.7 (C5′), 54.94 (CH3O), 54.92
Rf: 0.4 (10% MeOH in CH2Cl2, v/v); MALDI-HRMS m/z (CH3O). A minor impurity of chloroform at 79.1 ppm was
1
481.1399 ([M + Na]+, C26H22N2O6·Na+, Calc. 481.1370; H NMR identified in the 13C NMR.
(DMSO-d6): δ 11.28 (s, 1H, ex, NH), 8.42 (d, 1H, J = 7.8 Hz, Py),
8.29–8.31 (m, 2H, Py), 8.24–8.26 (d, 1H, J = 7.3 Hz, Py),
5′-O-(4,4′-Dimethoxytrityl)-2′-O-(pyren-4-yl)methyl-uridine (3Z)
8.18–8.22 (m, 3H, Py), 8.07 (t, 1H, J = 7.5 Hz, Py), 8.03 (dd, 1H, Nucleoside 2Z (150 mg, 0.33 mmol), DMTrCl (191 mg,
J = 7.8 Hz, 7.5 Hz, Py), 7.77 (d, 1H, J = 8.0 Hz, H6), 6.08 (d, 1H, 0.49 mmol) and DMAP (∼4 mg) in anhydrous pyridine (5 mL)
J = 5.4 Hz, H1′), 5.39–5.43 (m, 3H, 1 ex, 3′-OH, H5 and CH2Py), were mixed, reacted, worked up and purified as described
5.21–5.24 (d, 1H, J = 12.5 Hz, CH2Py), 5.09 (t, 1H, ex, 5′-OH), above to afford 3Z (200 mg, 80%) as a pale yellow foam. Rf:
4.29–4.33 (m, 1H, H3′), 4.20–4.23 (m, 1H, H2′), 3.97–4.00 0.6 (5% MeOH in CH2Cl2, v/v); MALDI-HRMS m/z 783.2695
(m, 1H, H4′), 3.63–3.68 (m, 1H, H5′), 3.57–3.62 (m, 1H, H5′); ([M
+
Na]+, C47H40N2O8·Na+, Calc. 783.2677); 1H NMR
13C NMR (DMSO-d6): δ 162.8, 150.5, 140.0 (C6), 132.8, 130.8, (DMSO-d6):36 δ 11.36 (s, 1H, ex, NH), 8.45 (d, 1H, J = 7.8 Hz,
130.5, 129.9, 129.0, 127.5 (Py), 127.0 (Py), 126.9 (Py), 126.3 (Py), Py), 8.17–8.33 (m, 6H, Py), 8.05 (t, 2H, H2Py, H7Py), 7.60 (d, 1H,
125.9 (Py), 125.3 (Py), 125.2 (Py), 124.1, 123.5, 121.7 (Py), 101.6 J = 8.2 Hz, H6), 7.30–7.32 (m, 2H, DMTr), 7.17–7.25 (m, 7H,
(C5), 86.0 (C1′), 85.4 (C4′), 80.7 (C2′), 70.2 (CH2Py), 68.4 (C3′), DMTr), 6.78–6.84 (m, 4H, DMTr), 6.07 (d, 1H, J = 4.1 Hz, H1′),
60.6 (C5′).
5.49 (d, 1H, J = 6.2 Hz, ex, 3′-OH), 5.44–5.46 (m, 1H, J =
12.7 Hz, CH2Py), 5.30–5.34 (d, 1H, J = 12.7 Hz, CH2Py), 5.08 (d,
1H, J = 8.2 Hz, H5), 4.38–4.42 (m, 1H, H3′), 4.26–4.28 (m, 1H,
H2′), 4.09–4.12 (m, 1H, H4′), 3.71 (s, 3H, CH3O), 3.69 (s, 3H,
General O5′-DMTr-protection protocol for the preparation of
3Y/3Z (description for ∼1 mmol scale)
The appropriate nucleoside 2 (specific quantities given below) CH3O), 3.30–3.34 (m, 1H, H5′ – partial overlap with H2O),
was co-evaporated twice with anhydrous pyridine and redis- 3.22–3.25 (m, 1H, H5′); 13C NMR (DMSO-d6): δ 162.8, 158.1,
solved in anhydrous pyridine. To this was added 4,4′- 158.0, 150.4, 144.4, 140.0 (C6), 135.3, 135.0, 132.7, 130.9,
dimethoxytritylchloride (DMTrCl) and catalytic N,N-dimethyl- 130.5, 129.9, 129.7 (DMTr), 129.6 (DMTr), 129.0, 127.8 (DMTr),
4-aminopyridine (DMAP), and the reaction mixture was stirred 127.6 (DMTr), 127.5 (Py), 127.1 (Py), 126.8 (Py), 126.7 (DMTr),
at rt under an argon atmosphere until TLC indicated complete 126.3 (C2Py), 126.0 (C7Py), 125.29 (Py), 125.25 (Py), 124.1, 123.5,
conversion (∼14 h). The reaction mixture was diluted with 121.7 (Py), 113.2 (DMTr), 113.1 (DMTr), 101.4 (C5), 86.9 (C1′),
CH2Cl2 (40 mL) and the organic phase was sequentially 85.9, 83.2 (C4′), 80.5 (C2′), 70.2 (CH2Py), 68.7 (C3′), 62.8 (C5′),
washed with water (2 × 30 mL) and sat. aq. NaHCO3 (2 × 55.0 (CH3O).
50 mL). The organic phase was evaporated to near dryness and
the resulting crude co-evaporated with abs. EtOH and toluene
(2 : 1, v/v, 3 × 3 mL) and purified by silica gel column chrom-
General O3′-phosphitylation protocol for the preparation of
4Y and 4Z (description for ∼0.25 mmol scale)
atography (0–5%, MeOH in CH2Cl2, v/v) to afford nucleoside 3 The appropriate nucleoside 3 (specific quantities given below)
(yield specified below).
was co-evaporated with anhydrous 1,2-dichloroethane and
redissolved in anhydrous CH2Cl2. To this was added N,N-diiso-
propylethylamine (DIPEA) and 2-cyanoethyl-N,N-diisopropyl-
5′-O-(4,4′-Dimethoxytrityl)-2′-O-(pyren-2-yl)methyl-uridine (3Y)
Nucleoside 2Y (500 mg, 1.09 mmol), DMTrCl (0.64 g, chlorophosphoramidite (PCl-reagent) and the reaction mixture
1.63 mmol) and DMAP (∼9 mg) in anhydrous pyridine (10 mL) was stirred at rt under an argon atmosphere until TLC
were mixed, reacted, worked up and purified as described indicated complete conversion (∼3 h), whereupon abs. EtOH
above to afford 3Y (0.57 g, 68%) as a pale yellow foam. Rf: (0.3 mL) and CH2Cl2 (5 mL) were sequentially added.
0.6 (5% MeOH in CH2Cl2, v/v); MALDI-HRMS m/z 783.2706 The organic phase was washed with sat. aq. NaHCO3 (2 mL),
([M
(DMSO-d6):35 δ 11.43 (s, 1H, ex, NH), 8.27–8.31 (m, 4H, H1Py, by silica gel column chromatography (40–70% EtOAc in
+
Na]+, C47H40N2O8·Na+, Calc. 783.2677); 1H NMR evaporated to near dryness, and the resulting residue purified
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petroleum ether, v/v) to afford phosphoramidite 4 (yield speci- the experiment. Quartz optical cells with a path length of
fied below).
1.0 cm were used. Thermal denaturation temperatures (Tm’s)
of duplexes (1.0 µM final concentration of each strand) were
measured on a Cary 100 UV/VIS spectrophotometer equipped
with a 12-cell Peltier temperature controller and determined as
3′-O-(2-Cyanoethoxy(diisopropylamino)phosphinyl))-5′-O-(4,4′-
dimethoxytrityl)-2′-O-(pyren-2-yl)methyl-uridine (4Y)
Nucleoside 3Y (145 mg, 0.19 mmol), DIPEA (136 µL, the maximum of the first derivative of the thermal denatura-
0.76 mmol) and PCl-reagent (85 μL, 0.38 mmol) in anhydrous tion curve (A260 vs. T) recorded in medium salt buffer (Tm
CH2Cl2 (5 mL) were mixed, reacted, worked up and purified as buffer: 100 mM NaCl, 0.1 mM EDTA, and pH 7.0 adjusted with
described above to afford phosphoramidite 4Y (170 mg, 93%) 10 mM Na2HPO4 and 5 mM Na2HPO4). The temperature of the
as a white foam. Rf: 0.8 (5% MeOH in CH2Cl2, v/v); MALDI- denaturation experiments ranged from at least 15 °C below Tm
HRMS m/z 983.3746 ([M + Na]+, C56H57N4O9P·Na+, Calc. to 20 °C above Tm (although not below 3 °C). A temperature
983.3755); 31P NMR (CDCl3): δ 150.5, 150.2.
ramp of 0.5 °C min−1 was used in all experiments. Reported
Tm-values are averages of two experiments within 1.0 °C.
3′-O-(2-Cyanoethoxy(diisopropylamino)phosphinyl))-5′-O-(4,4′-
dimethoxytrityl)-2′-O-(pyren-4-yl)methyl-uridine (4Z)
Protocol – determination of thermodynamic parameters
Nucleoside 3Z (190 mg, 0.25 mmol), DIPEA (178 µL, Thermodynamic parameters for duplex formation were deter-
0.99 mmol) and PCl-reagent (111 μL, 0.49 mmol) in anhydrous mined through baseline fitting of denaturation curves (van’t
CH2Cl2 (5 mL) were mixed, reacted, worked up and purified as Hoff analysis) using software provided with the UV/VIS spectro-
described above to afford phosphoramidite 4Z (220 mg, 92%) meter. Bimolecular reactions, two-state melting behavior, and
as a white foam. Rf: 0.8 (5% MeOH in CH2Cl2, v/v); MALDI- a heat capacity change of ΔCp = 0 upon hybridization were
HRMS m/z 983.3766 ([M + Na]+, C56H57N4O9P·Na+, Calc. assumed.30 A minimum of two experimental denaturation
983.3755); 31P NMR (CDCl3) δ 150.1, 150.0.
curves were each analyzed at least three times to minimize
errors arising from baseline choice. Averages and standard
deviations are listed.
Protocol – synthesis and purification of ONs
Synthesis of Y/Z-modified ONs was performed on an auto-
mated DNA synthesizer using 0.2 μmol scale succinyl linked
Protocol – absorption spectra
LCAA-CPG (long chain alkyl amine controlled pore glass) UV-vis absorption spectra (range: 300–400 nm) were recorded
columns with a pore size of 500 Å. Standard protocols for at 5 °C using the same samples and instrumentation as in the
incorporation of DNA phosphoramidites were used. A ∼50-fold thermal denaturation experiments.
molar excess of modified phosphoramidites in anhydrous
Protocol – steady-state fluorescence emission spectra
acetonitrile (at 0.05 M), along with extended oxidation (45 s)
and coupling times (activator: 0.01 M 4,5-dicyanoimidazole, Steady-state fluorescence emission spectra of ONs modified
15 min) was used during hand-couplings, which resulted in with pyrene-functionalized monomers X–Z and the corres-
stepwise coupling yields of 4Y and 4Z of ∼99% and ∼98%, ponding duplexes with complementary DNA/RNA targets, were
respectively. Cleavage from solid support and removal of pro- recorded in non-deoxygenated thermal denaturation buffer
tecting groups was accomplished upon treatment with 32% (each strand at 1.0 μM concentration) and obtained as an
aq. ammonia (55 °C, 24 h). The crude ONs were purified via average of five scans using an excitation wavelength of λex
=
ion-pair reverse phase HPLC (XTerra MS C18 column) using 350, 345 or 340 nm for X-, Y- or Z-modified ONs, excitation slit
0.05 M triethylammonium acetate–water/acetonitrile gradient, 5.0 nm, emission slit 2.5 nm and a scan speed of 600 nm
followed by detritylation (80% aq. AcOH, 20 min), and precipi- min−1. Experiments were determined at 5 °C to ascertain
tation (NaOAc–NaClO4–acetone, −18 °C for 12–16 h). The iden- maximal hybridization of probes to DNA/RNA targets (a stream
tity of synthesized ONs was established through MALDI-MS of nitrogen was blown in to the chamber to prevent
analysis (Table S1†) recorded in positive ions mode on a Quad- condensation).
rupole Time-Of-Flight Tandem Mass Spectrometer (Q-TOF Pre-
miere) equipped with a MALDI source (Waters Micromass
Protocol – molecular modeling
LTD, U.K) using anthranilic acid as a matrix, while purity The initial structures of the X/Y/Z-modified DNA duplexes were
(>85%) was verified by ion-pair reverse phase HPLC running in generated by building and modifying a standard B-type DNA
analytical mode.
duplex using MacroModel v9.8.37 The charge of the phospho-
diester backbone was neutralized with sodium ions, which
were placed 3.0 Å from non-bridging oxygen atoms and
Protocol – thermal denaturation studies
Concentrations of ONs were estimated using the following restrained to this distance by a force constant of 100 kJ mol−1
extinction coefficients for DNA (OD μmol−1):
G
(12.01), Å−2. A preliminary minimization was carried out using the
A (15.20), T (8.40), C (7.05); for RNA (OD μmol−1): G (13.70), Polak–Ribiere conjugate gradient method (convergence criteria
A (15.40), U (10.00), C (9.00); for pyrene (OD μmol−1): (22.4)31 0.1 kJ mol−1 Å−1), the AMBER94 force field38 with the
Strands were thoroughly mixed and denatured by heating to improved parambsc0 parameter set,39 and the implicit GB/SA
70–85 °C, followed by cooling to the starting temperature of continuum solvation model40 as implemented in MacroModel
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Paper
v9.8; all atoms were frozen except those in the O2′-substituent on X-ray film, which was developed using an X-Omatic 1000A
of the modified monomers. Non-bonded interactions were X-ray film developer (Kodak). The resulting bands were quanti-
treated with extended cut-offs (van der Waals 8.0 Å and electro- fied using a Fluor-S MultiImager (Bio-Rad, Hercules, CA)
statics 20.0 Å).
equipped with Quantity One software. Invasion efficiency was
The seed structure for X2:D4 was generated by taking pre- determined as the intensity ratio between the recognition
viously reported NMR data on X-modified DNA duplexes into complex band and the total lane. An average of three indepen-
account.31 Thus, distances between atoms for which NOE con- dent experiments is reported along with standard deviations.
tacts have been reported were constrained to 3.0–5.0 Å by a
force constant of 100 kJ mol−1 Å−2 (i.e., distances between (i)
Definition of zipper nomenclature
pyrene and H5 of the modified uridine, (ii) pyrene and H2/H8
The following nomenclature describes the relative arrange-
of the 3′-flanking adenosine, and (iii) pyrene and H6 of the
thymine opposite of the 3′-flanking adenosine). The structure
was minimized as described above with the following consider-
number of base pairs and has a positive value if a monomer is
ations: (i) all atoms were allowed to move freely during mini-
ment between two monomers positioned on opposing strands
in a duplex. The number n describes the distance measured in
shifted toward the 5′-side of its own strand relative to a second
reference monomer on the other strand. Conversely, n has a
negative value if a monomer is shifted toward the 3′-side of its
own strand relative to a second reference monomer on the
mization, (ii) sodium ions were restrained as above, and (iii)
hydrogen bonding of the outermost base pairs were restrained
by a force constant of 100 kJ mol−1 Å−2 ((C)2–O⋯HN–2(G), dis-
tance 1.99 Å, (C)N3⋯H–N1(G), distance 1.90 Å and (C)4–
NH⋯O–6(G), distance 1.69 Å). The resulting lowest energy
structure formed the basis for construction of the seed struc-
tures for all other studied duplexes. Pyrene moieties of the
other duplexes were manually adjusted to intercalated posi-
tions and the duplexes minimized as described for X2:D4.
Seed structures were then submitted to 5 ns of stochastic
dynamics (simulation temperature 300 K, time step 2.2 fs,
SHAKE all bonds to hydrogen) using the same constraints as
employed above (except for NOE constraints), during which
1000 structures were sampled and subsequently minimized.
The resulting structures were analyzed using MacroModel v9.8
and the 3DNA web server.41
other strand.
Acknowledgements
This study was supported by Award Number GM088697 from
the National Institute of General Medical Sciences, National
Institutes of Health; Awards IF13-001 and IF14-012 from the
Higher Education Research Council, Idaho State Board of Edu-
cation; The Office of Naval Research (N00014-10-1-0282);
INBRE Program, NIH grant no. P20 RR016454 (National Center
for Research Resources) and P20 GM103408 (National Institute
of General Medical Sciences); and Minitube of America. A.S.M.
is a Sapere Aude Postdoctoral Fellow (The Danish Council for
Independent Research). We thank Dr. Lee Deobald (EBI
Murdock Mass Spectrometry Center, Univ. Idaho) for assist-
ance with mass spectrometric analysis, Prof. Carolyn Bohach
(Food Science, Univ. Idaho) for access to gel documentation
stations, and Prof. Jean’ne Shreeve (Dept. Chemistry, Univ.
Idaho) for access to Parr reactors.
Protocol – electrophoretic mobility shift assay
This assay was performed, essentially as previously
described.13 Unmodified DNA hairpins DH1–DH7 were
obtained from commercial sources and used without further
purification. The DNA hairpins were 3′-DIG-labeled using the
2nd generation DIG Gel Shift Kit (Roche Applied Bioscience) as
recommended by the manufacturer. DIG-labeled ONs obtained
in this manner, were diluted and used without further purifi-
cation in the recognition experiments. Pre-annealed probes
(90 °C for 10 min, cooled to room temperature over 15 min)
and DIG-labeled DNA hairpins (34.4 nM) were mixed and incu-
bated in HEPES buffer (50 mM HEPES, 100 mM NaCl, 5 mM
MgCl2, 10% sucrose, 1.44 mM spermine tetrahydrochloride,
pH 7.2) for 15 h at ambient temperature (∼21 3 °C). The
reaction mixtures were then diluted with 6x DNA loading dye
(Fermentas) and loaded onto a 16% non-denaturing polyacryl-
amide gel. Electrophoresis was performed using a constant
voltage of 70 V for 2 h at ∼4 °C. Gels were blotted onto a posi-
tively charged nylon membrane (Roche Applied Bioscience)
using constant voltage with external cooling (100 V, ∼4 °C).
The membranes were exposed to anti-digoxigenin-AP Fab frag-
ments as recommend by the manufacturer of the DIG Gel Shift
Kit, transferred to a hybridization jacket, and incubated with
the substrate (CSPD) in detection buffer for 10 min at 37 °C.
The chemiluminescence of the formed product was captured
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