Synthesis and characterization of RNA containing a rigid and nonperturbing cytidine-derived spin label

Article abstract of DOI:10.1021/jo301227w

The nitroxide-containing nucleoside Cm is reported as the first rigid spin label for paramagnetic modification of RNA by solid-phase synthesis. The spin label is well accommodated in several RNA secondary structures as judged by its minor effect on the thermodynamic stability of hairpin and duplex RNA. Electron paramagnetic resonance (EPR) spectroscopic characterization of mono-, bi-, and trimolecular RNA structures shows that Cm will be applicable for advanced EPR studies to elucidate structural and dynamic aspects of folded RNA.

Full text of DOI:10.1021/jo301227w

Note
pubs.acs.org/joc
Synthesis and Characterization of RNA Containing a Rigid and
Nonperturbing Cytidine-Derived Spin Label
Claudia Hobartner,*,† Giuseppe Sicoli,‡ Falk Wachowius,† Dnyaneshwar B. Gophane,§
̈
and Snorri Th. Sigurdsson*,§
†Max Planck Research Group Nucleic Acid Chemistry and ‡Research Group Electron Paramagnetic Resonance, Max Planck Institute
for Biophysical Chemistry, Am Fassberg 11, 37077 Gottingen, Germany
̈
§Science Institute, University of Iceland, Dunhaga 3, 107 Reykjavik, Iceland
S
* Supporting Information
ABSTRACT: The nitroxide-containing nucleoside Çm is reported as the
first rigid spin label for paramagnetic modification of RNA by solid-phase
synthesis. The spin label is well accommodated in several RNA secondary
structures as judged by its minor effect on the thermodynamic stability of
hairpin and duplex RNA. Electron paramagnetic resonance (EPR)
spectroscopic characterization of mono-, bi-, and trimolecular RNA
structures shows that Çm will be applicable for advanced EPR studies to
elucidate structural and dynamic aspects of folded RNA.
flexible linkers and labels to structural constraints, rotational
freedom of the spin label can complicate the analysis of EPR
experiments.12 Although the number of reports on structural
investigations on RNA by EPR spectroscopy using established
spin labels is increasing,13−15 new EPR techniques and
instrumentation16 demand the development of new rigid spin
labels for RNA.17
itroxide spin labels have found widespread applications as
Nprobes for examination of both structure and dynamics of
biomolecules by EPR spectroscopy. In site-directed spin
labeling (SDSL), nitroxide probes are inserted at specific
positions in macromolecules, often through postsynthetic
labeling of prefunctionalized sites.1−4 For RNA, such function-
alized modifications include 2′-amino-modified pyrimidine
nucleotides,5,6 the phosphorothioate backbone,7,8 and 5-
iodopyrimidine or 2-iodoadenine.9,10 Alternatively, spin labels
are postsynthetically incorporated into RNA via convertible
nucleosides.11 This strategy yields spin-labeled nucleoside N4-
TEMPO-cytidine (CT) which contains the nitroxide label
attached at the exocyclic amino group of the nucleobase
(Figure 1). Postsynthetic modification strategies are usually tied
to the installation of flexible or semirigid tethers between the
attachment site and the nitroxide. Despite the adaptability of
Here, we report the 2′-O-methylribonucleoside C
spin label for paramagnetic labeling of RNA. Efficient
incorporation of Cm into RNA by solid-phase synthesis via
̧m as rigid
̧
compound 1 provides the first example of RNA spin labeling
using the phosphoramidite approach. The design of the spin
label Çm was inspired by the 2′-deoxyribonucleoside Ç (“C-
spin”, Figure 1), a rigid spin label for DNA in which the
nitroxide is fused to the nucleobase to form a phenoxazine
derivative.18−22
Çm was prepared from 2′-O-methyluridine (Um) in eight
steps (Scheme 1), starting with bromination of Um at position
5 to give 5-bromo-2′-O-methyluridine (2). Activation of 2 with
PPh3/CCl4, followed by coupling with the tetramethyl
isoindoline derivative 5 in the presence of DBU, which was
used to make Ç,
19 gave low yields of 6. Instead, the protected 5-
bromouridine derivative 3 was chlorinated at position 423 to
yield the dihalogenated nucleoside 4. Upon reaction with the
tetramethyl isoindoline derivative 5, the first nucleophilic
substitution occurred at the most reactive position of the
pyrimidine heterocycle. Treatment of compound 6 with
Figure 1. Nitroxide-containing cytidine derivatives and the phosphor-
amidite 1 for incorporation of Çm into RNA.
Received: June 23, 2012
Published: August 12, 2012
© 2012 American Chemical Society
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Scheme 1. Synthesis of Cm Phosphoramidite 1
Note
̧
potassium fluoride in ethanol yielded the ring-closed
phenoxazine derivative 7. The substitution reaction of 4 with
5 was preferably carried out in the presence of triethylamine,
which avoided the tedious removal of DBU before cyclization
to 7. Oxidation of compound 7 with hydrogen peroxide in the
presence of sodium tungstate gave the paramagnetic nucleoside
Çm. The 5′-hydroxyl group was protected as 4,4′-dimethoxy-
trityl ether using DMT-Cl and DMAP in pyridine. Subsequent
3′-phosphitylation in the presence of N,N-diisopropyl ammo-
nium tetrazolide and 2-cyanoethyl N,N,N′,N′-tetraisopropyldia-
midophosphite yielded the C
used for incorporation of C
solid-phase synthesis.
̧
m phosphoramidite 1, which was
m into RNA oligonucleotides by
Figure 2. (a) Anion exchange HPLC of crude and purified C
RNA 8. (b) CW-EPR spectra of single-stranded 8C
+9.
̧
m-labeled
m and duplex 8Çm
̧
̧
Tetramethyl-substituted nitroxyl radicals are moderately
sensitive to the conditions encountered during oxidation and
detritylation in the standard solid-phase synthesis cycle.10,20,24
Replacing the generally used iodine oxidation reagent with
tBuOOH and employing dichloroacetic acid instead of
trichloroacetic acid for detritylation have been shown to reduce
decomposition of nitroxides during DNA synthesis.20 We have
successfully adapted the tBuOOH oxidation conditions to
render the incorporation of spin-labeled phosphoramidite 1
compatible with RNA synthesis using 2′-O-TOM-protected
RNA phosphoramidites. After completion of the synthesis,
cleavage from solid support, deprotection of nucleobases and
phosphates was performed with methylamine, and the 2′-O-
TOM groups were removed by treatment with TBAF. The
RNA oligonucleotides were purified by anion exchange HPLC
and characterized by ESI-MS (Table S1 and Figure S1 in the
Supporting Information).
Analysis of the duplex conformation by CD spectroscopy
showed a slightly shifted and decreased maximum of the
ellipticity (Figure S3). This small difference compared to the
unmodified RNA is consistent with the spin label Çm being
well tolerated in an A-form helix, indicating that the extension
of the nucleobase is well accommodated in the major groove of
the double helix.
The new spin label Çm was then incorporated into a series of
mono- and bimolecular RNA structures (Figure 3), and its
properties were compared to the previously reported more
flexible RNA spin label CT.11 The spin labels Cm and CT, as
̧
well as the diamagnetic Cm, were individually incorporated
either at nucleotide C6 of the 20-mer RNA, giving the single-
labeled RNAs 10a, or simultaneously at C6 and C16, yielding
the double-labeled RNAs 10b. The complementary RNA 11
was modified at C7′ (primed numbers used for RNA 11),
resulting in the oligonucleotide 11a. RNAs 10 and 11 are
partially self-complementary and can therefore fold into
hairpins (with four unpaired nucleotides at the 3′ or the 5′
terminus). The unpaired overhangs of 10 and 11 can hybridize
to those of hairpins 12 and 13, respectively, which results in the
formation of bimolecular dumbbell structures (10+12 and
11+13). Using different combinations of RNAs 10, 10a, 10b,
11, and 11a, four spin-labeled duplex samples were studied: two
samples with a single label at position C6 or C7′, and two
double-labeled duplexes with labels at nucleotides C6 and C16
and C6 and C7′, respectively.
The efficient incorporation of C
demonstrated by synthesis of the 14-mer oligoribonucleotide
8Cm (Figure 2a). Thermodynamic analysis by UV melting
experiments determined the influence of Cm on duplex
stability. The 14-mer Cm-labeled duplex (8Cm+9) showed a
̧m into RNA was first
̧
̧
̧
̧
decrease in melting temperature by only 1.5−2.5 °C at various
concentrations between 1 and 30 μM, compared to the
unmodified (8C+9) or 2′-O-methylcytidine (Cm)-containing
duplexes (8Cm+9) (Table 1 and Figure S2). On the basis of
van’t Hoff analysis of concentration-dependent melting curves,
the difference in Tm translates into a ΔΔG298 of 4.0 kcal/mol
compared to the unmodified RNA (Figure S2 and Table S2).
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Note
Table 1. UV Melting Analysis
d
a
C
Cm
C
̧
m
a
CT
Tm
nd
a
a
b
c
b
RNA
8+9
Tm
Tm
ΔTm
+0.9
+1.7
Tm
ΔTm
−1.6/−2.5
−0.4/−2.1
ΔTm
D
D
D
D
D
H
H
H
75.1
82.2
82.2
82.2
82.2
88.2
88.2
76.4
76.0
83.9
85.5
82.5
83.1
88.5
89.5
73.5
81.8
80.1
78.3
84.0
88.8
89.7
73.4
nd
−7.1
10a+11
10b+11
10a+11a
10+11a
10a
75.1
69.4
nd
+3.3 (+1.7)
+0.3 (+0.2)
+0.9
−2.1/−5.4 (−1.1/−2.7)
−3.9/−4.2 (−1.9/−2.1)
+1.7/+0.9
−12.8 (−6.4)
nd
nd
nd
+0.3
+0.6/+0.3
75.4
62.7
nd
−12.8
10b
+1.3 (+0.7)
+1.4
+1.5/+0.2 (+0.8/0.1)
−3.0/−4.4
−25.5 (−12.8)
11a
77.8
b
nd
a
c
Melting temperature in °C ( 0.5 °C). ΔTm to unmodified RNA. ΔTm to unmodified/Cm-modified RNA (ΔTm per modification is given in
d
parentheses). Data from ref 11. D, duplex; H, hairpin; nd, not determined.
spectrum is characteristic for the slow-motion regime of
nitroxyl radicals in spin-labeled proteins27 and oligonucleotides
and compares well with the reported label C
DNA sequence (Figure S6).19 This observation validates the
rigidity of Cm and its restricted motion in an RNA duplex, due
̧ in an analogous
̧
to base pairing and stacking interactions. In contrast, the line
shape of the CW-EPR spectrum of the single-stranded RNA
8C
and unstructured single strand containing a rigid spin label.20
The CW-EPR spectrum of the Cm-labeled hairpin 10aC
exhibited three resolved hyperfine lines (Figure 4A), with a
̧m reflects intermediate mobility, as expected for a flexible
̧
̧m
Figure 3. Sequences and secondary structures of RNAs. Individually
modified cytidines are indicated in red. Modified (Cm-, CT-, and C
̧m-
containing) and unmodified RNA oligonucleotides are in black and
green, respectively.
The influence of the spin labels on the stability of the hairpin
and duplex structures was assayed by UV melting experiments
(Table 1, Figures S4 and S5). Only minor changes in Tm of less
than +1 °C per spin label were observed for the hairpins
10aC
at the loop closing and the terminal base pair of the hairpin
stem. Placing Cm at an internal base pair in the stem of 11a
resulted in a moderate destabilization by 3 °C. In all of the
bimolecular duplex structures, the change in Tm caused by C
was less than 2 °C per modification relative to the unmodified
RNA. The slightly different effects of Cm at different positions
̧m and 10bÇm, indicating that Çm is well accommodated
̧
̧
m
̧
of the same duplex sequence most likely reflect flanking-
sequence dependence, previously observed with phenoxazine
derivatives in DNA.20,25
By comparison of the Tm values with the analogous Cm-
modified RNAs, the effect of the extended tetracyclic
nucleobase was extracted. As expected, the 2′-OMe group has
Figure 4. CW-EPR spectra of CT and C
different secondary structure contexts.
̧m spin-labeled RNAs in
a positive effect on the overall stability of the C
̧
m-modified
significantly broader line shape than the CT-labeled hairpin
RNA 10aCT (Figure 4B). The Cm- and CT-labeled duplexes
RNA, likely due to preorganization of the ribose in the north-
type C3′-endo sugar conformation as usually found in RNA
duplexes.26 Thus, the destabilizing effect of the nucleobase
modification is counter-balanced by the 2′-ribose modification.
̧
(Figure 4C,D) show additional broadening, due to their
increased size and associated slower rotational correlation
times. The observed hyperfine splitting of 177 MHz for the
The comparison of the new label C
̧
m with the more flexible
Çm-labeled duplex (Figure 4C) is typical for rigid nitro-
RNA label CT11 reveals considerably less destabilization by C
(Table 1). This demonstrates that Cm is better tolerated than
CT in the investigated hairpin and duplex structures.
̧
m
xides19,27 and characteristic for the slow-motion regime of
nitroxide labels with correlation times in the range of 10 ns.28
In contrast, such large splitting was not observed for the CT-
labeled duplex (Figure 4D); the detected line shape reflects a
correlation time on the order of 5 ns.28 Analogous effects were
observed for hairpin and duplex structures containing RNAs
̧
The spin-labeled RNA structures were then examined by
EPR spectroscopy. The continuous wave (CW)-EPR spectra of
the single-stranded RNA 8C
shown in Figure 2b. The line shape of the duplex CW-EPR
̧m and the duplex 8Çm+9 are
10bÇm and 11aÇm (Figure S7A−D). A summary of the
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Note
characteristic hyperfine splitting values 2Azz (separation
between low- and high-field resonances) and the widths of
the central lines ΔH0 are reported in Table S3 and Figure S8. A
local environment of the spin probe. In addition, C
̧
m will be
highly valuable for orientation selection experiments that allow
determination of the relative orientation of spin labels within
biomolecules. This will enable the structural and dynamic
characterization of larger multihelix RNAs, which are of current
interest as regulators of gene expression and components of
RNA maturation machineries.
comparison of the CW-EPR spectra of Ç
m and CT-labeled
hairpin and duplex RNAs at 22 and 10 °C (Figure S9) suggests
that the new label Cm may be used to detect temperature-
̧
dependent changes in local base pair dynamics that are not
reported by CT.
EXPERIMENTAL SECTION
To further examine RNA secondary structures with Çm, we
explored the formation of bimolecular dumbbell structures,
which contain the spin-labeled hairpins 10a and 11a. As
■
General. Thin layer chromatography (TLC) was carried out using
glass plates precoated with silica gel (0.25 mm, F-254). Compounds
were visualized by UV light and staining with p-anisaldehyde. Flash
column chromatography was performed using ultrapure flash silica gel
(230−400 mesh size, 60 Å). Dichloromethane and pyridine were
freshly distilled from calcium hydride prior to use. Anhydrous
triethylamine, n-hexane, and ethyl acetate were used without further
purification. All moisture- and air-sensitive reactions were carried out
in oven-dried glassware under an inert argon atmosphere. NMR
spectra were recorded on a 400 MHz spectrometer. Commercial grade
CDCl3 was passed over basic alumina shortly before use with tritylated
expected, the CW-EPR line shapes of the C
dumbbells 10aCm+12 and 11aCm+13 (Figure 4E, Figure
S7E) reflect again the strongly restricted mobility of the spin
label C
m, whereas the spectrum of the CT-labeled dumbbell
(Figure 4F) is insignificantly different from the hairpin
spectrum. The visible spectral broadening for Cm-labeled
̧m-labeled
̧
̧
̧
̧
dumbbells (compared to hairpins) is consistent with a
decreased rotational correlation time due to the extended
length of the base-paired stem upon hybridization of the
complementary overhangs. We have used this effect to
distinguish interacting from noninteracting RNA sequences.
For example, the identical single-stranded overhangs of hairpins
1
compounds. H NMR chemical shifts are reported in reference to
undeuterated residual solvent in CDCl3 (7.26 ppm), DMSO-d6 (2.50
ppm), and MeOH-d4 (3.31 and 4.84 ppm). 13C NMR chemical shifts
are reported in reference to solvent signal (CDCl3 (77.16 ppm),
DMSO-d6 (39.43 ppm), and MeOH-d4 (49.05 ppm)). 31P NMR
chemical shifts are reported relative to 85% H3PO4 as an external
standard. NMR spectra of nitroxide-containing compounds show
significant broadening, sometimes to the extent that some nuclei are
10aÇm and 13 cannot interact via standard Watson−Crick
base pairs. Therefore, upon mixing 10a and 13, the
monomolecular hairpin structures remain intact and display
an unchanged CW-EPR spectrum (i.e., no dumbbell is formed,
Figure S7F). In an additional set of experiments, we explored
trimolecular RNA structures, in which RNA 11a was hybridized
with 2 equiv of short unlabeled RNAs (Figure S10). These
trimolecular structures are essentially duplexes with gap(s). A
duplex containing a four-nucleotide gap had considerably
higher mobility than a duplex containing a single-nucleotide
1
not observed in the spectra. For the same reason, integration of H
NMR spectra of nitroxides is not reported. Mass spectrometric
analyses of all organic compounds were performed on an HR-ESI-MS
(MicroTof-Q) in positive ion mode.
5-Bromo-2′-O-methyluridine (2). To a suspension of 2′-O-
methyluridine (Um, 2.00 g, 7.74 mmol) in 1,2-dimethoxyethane (43
mL) was added a solution of NaN3 (2.51 g, 38.7 mmol) in water (3.2
mL). [Note: NaN3 is toxic and hazardous to the environment, and
bulk quantities pose an explosion hazard.] The resulting mixture was
stirred for 15 min at 22 °C and treated with N-bromosuccinimide
(1.79 g, 10.0 mmol). The reaction mixture was stirred for 24 h at 22
°C. The solvent was removed in vacuo and the residue passed through
a short silica gel column (CH2Cl2/MeOH; 100:0 to 95:5) to yield 5-
bromo-2′-O-methyluridine (2) as a white solid (2.30 g, 88% yield): mp
gap, demonstrating the sensitivity of the Çm probe for
evaluating the flexibility of hinges/junctions between two
helical regions.
The C
for example, to explore RNA structures by measuring interspin
distances. Additionally, the rigid spin label Cm provides strong
̧m label is also well suited for pulsed EPR experiments,
̧
1
223−225 °C; H NMR (DMSO-d6) δ 3.39 (s, 3H), 3.56−3.61 (m,
orientation selection in the distance measurements. This allows
for determination of orientations even at low EPR frequency (9
1H), 3.68−3.73 (m, 1H), 4.10−4.13 (dd, J = 5.3 Hz, 11.1 Hz, 1H),
5.12−5.14 (d, J = 6.3 Hz, 1H), 5.32−5.34 (t, J = 4.6 Hz, 9.2 Hz, 1H),
5.79 (d, J = 3.8 Hz, 1H), 8.53 (s, 1H), 11.83 (br s, 1H); 13C NMR
(DMSO-d6) 57.5, 59.5, 67.7, 82.9, 84.7, 86.6, 95.7, 140.0, 149.6, 159.1;
HR-ESI-MS m/z calcd for C10H13BrN2O6 (M + Na)+ 358.9849, found
358.9841.
GHz), as already shown for Ç
-labeled DNA duplexes,12,22 but it
also complicates the distance analysis. Pulsed electron double
resonance (PELDOR) experiments with RNA samples
containing two C
in agreement with estimations for typical A-form RNA helices
(Figure S11). An extensive high-field pulsed EPR study of Cm-
̧m labels provided distance results that are
3′,5′-Diacetyl-5-bromo-2′-O-methyluridine (3). Acetic anhy-
dride (4.18 mL, 41.1 mmol) was added to a solution of 2 (2.30 g, 6.84
mmol) in dry pyridine (5.52 mL) at 0 °C. The reaction mixture was
stirred at 22 °C for 16 h, and the solvent was removed in vacuo. The
residue was dissolved in CH2Cl2 (70 mL) and washed with water (3 ×
70 mL). The organic layer was dried over anhydrous Na2SO4, filtered,
and concentrated to give the crude product, which was purified by
column chromatography (CH2Cl2/MeOH; 100:0 to 97:3) to yield
compound 3 as a white solid (2.50 g, 87% yield): mp 162−164 °C; 1H
NMR (CDCl3) δ 2.15 (s, 3H), 2.23 (s, 3H), 3.51 (s, 3H), 4.03−4.05
(dd, J = 2.5 Hz, 5.1 Hz, 1H), 4.38−4.43 (m, 3H), 4.94 (dd, J = 7.2, 5.3
Hz, 1H), 5.93 (d, J = 2.5 Hz, 1H), 7.97 (s, 1H), 9.29 (br s, 1H); 13C
NMR (CDCl3) δ 20.6, 21.1, 59.1, 61.8, 69.3, 79.3, 81.9, 88.5, 97.2,
138.5, 149.3, 158.6, 170.1, 170.2; HR-ESI-MS m/z calcd for
C14H17BrN2O8 (M + Na)+ 443.0060, found 443.0068.
̧
labeled RNA for distance and orientation measurements will be
reported in due course.
In summary, we have synthesized nucleoside Çm containing
a rigid nitroxide spin label, incorporated it into different RNA
structural contexts by solid-phase synthesis, and analyzed the
spin-labeled RNA by UV, CD, and EPR spectroscopy. In
several aspects, C
label CT and other previously reported spin probes for RNA.
The new label Cm was well tolerated in A-form helices, as
judged by its small influence on the thermodynamic stability of
labeled RNAs. By CW-EPR spectroscopy, Cm reported on the
̧m compared favorably with the more flexible
̧
̧
local environment of the labeling site and provided information
on the global RNA structure (hairpin versus duplex or
dumbbell). The rigid RNA label will be applicable not only
to study intermolecular interactions but will also prove useful
for sensing intramolecular refolding events or changes in the
Compound 4. A solution of compound 3 (0.50 g, 1.19 mmol) and
PPh3 (0.78 g, 2.97 mmol), in a mixture of CH2Cl2 and CCl4 (7 + 7
mL), was refluxed for 2.5 h. The solvent was removed in vacuo, and
the residue was purified by silica gel flash column chromatography
using a gradient elution (EtOAc/CH2Cl2; 5:95 to 15:85) to yield
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Note
1
compound 4 as a white solid (0.31 g, 60% yield): mp 66−68 °C; H
NMR (CDCl3) δ 2.14 (s, 3H), 2.23 (s, 3H), 3.63 (s, 3H), 4.12−4.14
(d, J = 4.9 Hz, 1H), 4.42−4.55 (m, 3H), 4.75−4.78 (dd, J = 4.5 Hz, 9.5
Hz, 1H), 5.89 (s, 1H), 8.39 (s, 1H); 13C NMR (CDCl3) δ 20.5, 21.1,
59.2, 61.0, 68.5, 79.1, 81.5, 90.1, 97.5, 143.7, 151.4, 166.1, 170.0, 170.1;
HR-ESI-MS m/z calcd for C14H16BrClN2O7 (M + Na)+ 460.9722,
found 460.9730.
152.09; HR-ESI-MS m/z calcd C52H62N6O10P (M + Na)+ 984.4157,
found 984.4191.
RNA Synthesis, Purification, and Characterization. RNA
solid-phase syntheses were performed on a Pharmacia Gene Assembler
Plus, using polystyrene custom primer support from GE Healthcare.
2′-O-TOM-protected ribonucleoside phosphoramidites were used for
all unmodified RNA nucleotide positions. CT-modified RNA was
prepared as previously described using a convertible O4-(4-
Compound 6. A solution of compounds 519 (87 mg, 0.42 mmol)
and 4 (223 mg, 0.51 mmol) in CH2Cl2 (10 mL) was treated with Et3N
(55.5 mg, 0.55 mmol) at 22 °C. After 16 h, the solvent was removed in
vacuo and the crude product was used directly for the next reaction
without further purification.
chlorophenyl)uridine phosphoramidite.11
synthesized using phosphoramidite 1. Since the C
Ç
m-modified RNA was
̧
m phosphoramidite
1 had limited solubility in acetonitrile, it was dissolved at a
concentration of 100 mM in 1,2-dichloroethane. The coupling time
was set to 3 min using 250 mM benzylthiotetrazole as a coupling
agent. Oxidation was performed with 1 M tert-butylhydroperoxide in
Compound 7. KF (245 mg, 4.22 mmol) was added to a solution of
compound 6 (257 mg, 0.42 mmol crude from previous step) in
absolute ethanol (32 mL). After refluxing for 5 days, the solution was
filtered and the solvent was removed in vacuo. The product was
purified by silica gel flash column chromatography (CH2Cl2/MeOH;
100:0 to 65:35) to yield 7 as dark yellow solid (93 mg, 50% yield): mp
toluene, inspired by recent reports on the stability of Ç under these
conditions.20 Capping and detritylation were performed under
standard conditions for RNA synthesis with 2′-O-TOM phosphor-
amidites. The Çm-containing RNA oligonucleotides were deprotected
1
(decomp.) 160−162 °C; H NMR (MeOH-d4) δ 1.44 (d, J = 1.2 Hz,
by treatment with 8 M methylamine in ethanol/H2O 1/1 at 37 °C for
5−6 h, followed by 2′-O-TOM deprotection with 1 M tetrabuty-
lammonium fluoride (TBAF) in THF at 25 °C for 12−16 h. After
desalting on a Sephadex G10 column, the quality of the crude product
was checked by analytical anion exchange HPLC on a Dionex
DNAPac PA200 column, 4.6 × 250 mm, flow rate 1 mL/min, 80 °C,
buffer A: 25 mM Tris·HCl pH 8.0, 6 M urea; buffer B: buffer A + 0.5
M NaClO4, linear gradient of B in A, with a slope of 4% per column
volume. RNA oligonucleotides were purified by semipreparative anion
exchange HPLC under denaturing conditions on a Dionex DNAPac
PA100 column, 9 × 250 mm, flow rate 2 mL/min, 80 °C, buffers A
and B as for analytical column. Detection was by UV absorbance at
280 nm. Fractions containing full-length RNA were collected and
desalted on SepPak cartridges (Waters). RNA concentration was
determined by UV absorbance at 260 nm, and the product identity was
confirmed by ESI-MS (Table S1). RNA samples were stored as
aqueous solutions at −20 °C.
12H), 3.57 (s, 3H), 3.74−3.84 (m, 2H), 3.91−3.97 (m, 2H), 4.21−
4.24 (dd, J = 6.8 Hz, 5.17 Hz, 1H), 5.89 (d, J = 2.6 Hz), 6.60−6.61 (d,
J = 4.1 Hz, 2H), 7.86 (s, 1H); 13C NMR (MeOH-d4) δ 31.0, 31.1,
58.9, 61.1, 64.9, 64.9, 69.3, 85.5, 89.6, 109.6, 110.7, 123.7, 127.8, 129.6,
143.7, 144.1, 144.6, 155.9, 156.3; HR-ESI-MS m/z calcd for
C22H28N4O6 (M + H)+ 445.2082, found 445.2093.
Rigid Spin Label Çm. A solution of 7 (90 mg, 0.20 mmol) in
MeOH (5.5 mL), containing NaHCO3 (17 mg, 0.20 mmol), was
treated dropwise with H2O2 (70% w/v, 1.11 g/mL, 0.06 mL, 1.42
mmol). After 5 min, Na2WO4 (10 mg, 0.03 mmol) was added and the
resulting mixture was stirred for 30 h at 22 °C. The salts were filtered
and discarded, the filtrate was concentrated in vacuo, and the residue
was purified by flash column chromatography (CH2Cl2/MeOH; 100:0
to 85:15) to yield Çm as a yellow solid (47 mg, 51% yield): mp
(decomp.) 230−232 °C; 1H NMR (DMSO-d6) δ 2.08 (br s), 3.41 (br
s), 3.70 (br s), 3.85 (br s), 4.13 (br s), 5.09 (br s), 5.26 (br s), 5.88 (br
s), 7.52 (br s), 8.54 (s), 10.72 (br s); 13C NMR (DMSO-d6) δ 30.5,
57.3, 59.6, 67.5, 82.8, 84.0, 86.0, 86.2, 95.4, 125.2, 139.6, 149.2, 151.2,
151.4, 158.6; HR-ESI-MS m/z calcd for C22H27N4O7 (M + Na)+
482.1772, found 482.1785.
Melting curves were measured in 10 mM potassium phosphate
buffer, pH 7.0, 150 mM NaCl, on a Cary 100 UV spectrophotometer
(Varian Inc.) equipped with a multiple cell holder and a Peltier
temperature-control device. RNA sample concentration was from 2 to
40 μM. Temperature-dependent changes in UV absorbance were
measured at 250, 260, 270, and 280 nm, with a heating/cooling rate of
0.7 °C/min. Two full heating and cooling cycles (4 ramps) were
collected, and all melting transitions were fully reversible and
reproducible. Thermodynamic parameters for the duplices 8+9 were
obtained from concentration-dependent melting curves by analysis of
ln(cT) versus 1/Tm.
5′-(4,4′-Dimethoxytrityl) Çm. Spin-labeled nucleoside Çm (20.0
mg, 0.04 mmol), DMTCl (22.1 mg, 0.06 mmol), and DMAP (0.6 mg,
0.004 mmol) were added to a round-bottom flask and kept under
vacuum for 16 h. Anhydrous pyridine (1 mL) was added, and the
resulting solution was stirred for 4 h, after which MeOH (100 μL) was
added and the solvent removed in vacuo. The residue was purified by
column chromatography (Et3N/CH2Cl2/MeOH; 1:99:0 to 1:95:4) to
yield tritylated C
̧
m as a yellow solid (20 mg, 60% yield): mp
CD spectra were recorded on a Chirascan CD spectrophotometer
(Applied Photophysics) at an RNA duplex concentration of 10 or 40
μM in 10 mM potassium phosphate buffer, pH 7.0, 150 mM NaCl, at
25 °C, with 0.5 nm step size and 2 s/data point. Data were collected in
three repetitions and averaged. CD spectra are depicted in Supporting
Information Figures S1 and S3.
(decomp.) 150−152 °C; 1H NMR (DMSO-d6) δ 3.45 (br s), 3.74 (br
s), 3.96 (br s), 4.25 (br s), 5.19 (br s), 5.83 (br s), 6.93 (br s), 7.33−
7.44 (br s), 10.65 (br s); 13C NMR (DMSO-d6) δ 57.6, 62.1, 68.1,
82.0, 82.7, 85.6, 86.9, 113.0, 113.1, 126.6, 127.6, 127.6, 127.7, 127.8,
129.4, 129.5, 135.1, 135.4, 144.2, 157.8; HR-ESI-MS m/z calcd
C43H45N4O9 (M + Na)+ 784.3079, found 784.3083.
EPR Experiments. Aliquots of RNA samples were lyophilized and
dissolved in buffer. The buffer composition was 10 mM potassium
phosphate buffer, pH 7.0 containing 150 mM NaCl. The final
concentration of the spin-labeled RNA was 10−25 μM. For
bimolecular samples with only one spin-labeled strand, the unmodified
complementary strand was used in 1.5-fold excess; for the trimolecular
constructs, the short unmodified strand (14 or 15) was in 10-fold
excess. For the duplex sample containing two single-labeled strands,
equimolar amounts of 10a+11a were used. All samples were heated to
90 °C for 3 min and slowly cooled to 22 °C. The EPR samples (15
μL) were placed in quartz capillaries sealed at one end. CW-EPR
spectra were recorded at 23 °C at X-band (9 GHz) over 160 G on a
Bruker Elexsys 500 spectrometer fitted with a high-sensitivity
resonator using 20 mW incident microwave power and 1 G field
modulation amplitude at 100 kHz modulation frequency. From the
CW-EPR spectra, the peak-to-peak line width of the central line (ΔH0,
in mT) and the separation between the resonances of the low- and
high-field lines (2Azz, in mT) were extracted (Supporting Table S3)
C
̧
m Phosphoramidite (1). Diisopropyl ammonium tetrazolide
(24 mg, 0.14 mmol) and tritylated Cm (70.0 mg, 0.09 mmol) were
̧
dissolved in pyridine, the pyridine was removed in vacuo, and the
residue was kept under vacuum for 16 h. CH2Cl2 (4 mL) was added,
along with 2-cyanoethyl N,N,N′,N′-tetraisopropyldiamidophosphite
(88 μL, 0.28 mmol), and the resulting solution was stirred for 5 h at 22
°C. CH2Cl2 (10 mL) was added, and the organic phase was washed
with saturated aqueous NaHCO3 (3 × 15 mL) and saturated aqueous
NaCl (3 × 15 mL), dried over Na2SO4, and concentrated in vacuo.
The residue was dissolved in a minimum amount of diethyl ether (7−9
mL), followed by a slow addition of hexane (40−50 mL) at 22 °C.
The solvent was decanted and discarded. Finally, the compound was
purified by column chromatography using neutral silica gel (EtOAc) to
yield 1 as a yellow solid (40 mg, 46% yield): mp (decomp.) 130−132
1
°C; H NMR (CDCl3) δ 1.24−1.63 (br m), 2.82 (br s), 3.04 (br s),
4.15 (br s), 4.50 (br s), 4.64 (br s), 4.87 (br s), 5.10 (br s), 5.62 (br s),
6.42 (br s), 7.26 (br s), 7.82 (br s); 31P NMR (CDCl3) δ 150.59,
7753
dx.doi.org/10.1021/jo301227w | J. Org. Chem. 2012, 77, 7749−7754

The Journal of Organic Chemistry
Note
and plotted as a function of the number of base pairs (Supporting
Figure S6).
(20) Cekan, P.; Smith, A. L.; Barhate, N.; Robinson, B. H.;
Sigurdsson, S. T. Nucleic Acids Res. 2008, 36, 5946.
(21) Cekan, P.; Jonsson, E. O.; Sigurdsson, S. T. Nucleic Acids Res.
2009, 37, 3990.
(22) Marko, A.; Denysenkov, V.; Margraf, D.; Cekan, P.; Schiemann,
O.; Sigurdsson, S. T.; Prisner, T. F. J. Am. Chem. Soc. 2011, 133,
13375.
(23) Denapoli, L.; Messere, A.; Montesarchio, D.; Piccialli, G.;
Santacroce, C. Bioorg. Med. Chem. Lett. 1992, 2, 315.
(24) Gannett, P. M.; Darian, E.; Powell, J.; Johnson, E. M., II;
Mundoma, C.; Greenbaum, N. L.; Ramsey, C. M.; Dalal, N. S.; Budil,
D. E. Nucleic Acids Res. 2002, 30, 5328.
ASSOCIATED CONTENT
■
S
* Supporting Information
List of abbreviations, Supporting Tables S1−S3, Supporting
Figures S1−S11, copies of 1H and 13C NMR spectra for
compounds 2−7, Çm, and DMT Ç
m, 1H and 31P NMR spectra
for phosphoramidite 1. This material is available free of charge
via the Internet at http://pubs.acs.org.
(25) Sandin, P.; Borjesson, K.; Li, H.; Martensson, J.; Brown, T.;
̈
̊
AUTHOR INFORMATION
Corresponding Author
*E-mail: claudia.hoebartner@mpibpc.mpg.de, snorrisi@hi.is.
Notes
■
Wilhelmsson, L. M.; Albinsson, B. Nucleic Acids Res. 2008, 36, 157.
(26) Egli, M.; Pallan, P. S. Annu. Rev. Biophys. Biomol. Struct. 2007,
36, 281.
(27) Hubbell, W. L.; Cafiso, D. S.; Altenbach, C. Nat. Struct. Biol.
2000, 7, 735.
The authors declare no competing financial interest.
(28) Stoll, S.; Schweiger, A. J. Magn. Reson. 2006, 178, 42.
ACKNOWLEDGMENTS
■
Prof. Marina Bennati (MPIbpc, Research Group Electron
Paramagnetic Resonance) is gratefully acknowledged for
stimulating discussion and access to EPR instruments. We
thank Uwe Pleßmann and Prof. Henning Urlaub (MPIbpc,
Facility for mass spectrometry) for recording ESI-MS spectra of
RNA oligonucleotides, and Brigitta Angerstein for help with
EPR sample preparation. This work was supported by the Max
Planck Society and the Icelandic Research Fund (090026023).
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