Probing RNA dynamics via longitudinal exchange and CPMG relaxation dispersion NMR spectroscopy using a sensitive 13C-methyl label

Article abstract of DOI:10.1093/nar/gkq1361

The refolding kinetics of bistable RNA sequences were studied in unperturbed equilibrium via ¹³C exchange NMR spectroscopy. For this purpose a straightforward labeling technique was elaborated using a 2′-¹³C-methoxy uridine modification, which was prepared by a two-step synthesis and introduced into RNA using standard protocols. Using ¹³C longitudinal exchange NMR spectroscopy the refolding kinetics of a 20nt bistable RNA were characterized at temperatures between 298 and 310K, yielding the enthalpy and entropy differences between the conformers at equilibrium and the activation energy of the refolding process. The kinetics of a more stable 32nt bistable RNA could be analyzed by the same approach at elevated temperatures, i.e. at 314 and 316K. Finally, the dynamics of a multi-stable RNA able to fold into two hairpin-and a pseudo-knotted conformation was studied by ¹³C relaxation dispersion NMR spectroscopy.

Full text of DOI:10.1093/nar/gkq1361

4340–4351 Nucleic Acids Research, 2011, Vol. 39, No. 10
doi:10.1093/nar/gkq1361
Published online 19 January 2011
Probing RNA dynamics via longitudinal exchange
and CPMG relaxation dispersion NMR spectroscopy
13
using a sensitive C-methyl label
Karin Kloiber1,2, Romana Spitzer1, Martin Tollinger1, Robert Konrat2 and
Christoph Kreutz1,*
2
1Institute of Organic Chemistry, Leopold Franzens University, Innrain 52a, 6020 Innsbruck and Institute of
Computational and Structural Biology, University of Vienna, Max F. Perutz Laboratories, Campus Vienna
Biocenter 5, 1030 Vienna, Austria
Received October 8, 2010; Revised December 23, 2010; Accepted December 30, 2010
ABSTRACT
certain gene. Closely related to this new species are RNA
based thermometers that switch between distinct folded
states depending on temperature, which subsequently
yields specific gene expression patterns in response to
temperature changes (1–6).
These non-coding ribonucleic acids illustrate a common
theme concerning their regulatory function: the existence
The refolding kinetics of bistable RNA sequences
were studied in unperturbed equilibrium via 13C
exchange NMR spectroscopy. For this purpose a
straightforward labeling technique was elaborated
using a 20-13C-methoxy uridine modification, which
was prepared by
a
two-step synthesis and
of distinct folded states of RNA that fulfill different tasks
(7–10). Frequently, RNA conformational landscapes
comprise several stable structures with only marginal
free energy differences. It follows that such RNA se-
quences can exist in multiple conformations at the same
time. Interconversions between different secondary struc-
tures represent a common regulatory mechanism for the
modulation of RNA function and constitute the basic
steps of large-scale conformational transitions.
Therefore, the thermodynamics and dynamics of such
processes deserve a detailed investigation.
For the purpose of characterizing RNA refolding
processes, Hobartner and Micura (8) introduced so-called
bistable RNAs, i.e. short sequences which populate two or
more conformational substates. These small systems
permit a detailed investigation of the dynamic properties
of isolated secondary structure elements. In this study we
chose bistable RNAs as appropriate model systems for the
introduction and the benchmarking of the presented
approach.
NMR spectroscopy has proven to be very well suited
for monitoring the exchange between distinct folded states
of RNA. The kinetics of various bistable RNAs have been
determined by applying an experimentally demanding
real-time NMR approach that relies on trapping a
certain conformation by the introduction of a photolabile
protection group. Subsequent laser-induced release of the
protection group initiates the return to the equilibrium
population of the respective bistable RNA, which is
introduced into RNA using standard protocols.
Using 13C longitudinal exchange NMR spectroscopy
the refolding kinetics of a 20 nt bistable RNA were
characterized at temperatures between 298 and
310 K, yielding the enthalpy and entropy differences
between the conformers at equilibrium and the
activation energy of the refolding process. The
kinetics of a more stable 32 nt bistable RNA could
be analyzed by the same approach at elevated tem-
peratures, i.e. at 314 and 316 K. Finally, the dynamics
of a multi-stable RNA able to fold into two hairpin-
and a pseudo-knotted conformation was studied by
13C relaxation dispersion NMR spectroscopy.
INTRODUCTION
It is only recently that RNA biology has gained a lot of
attention, due to the discovery that RNA fulfills a variety
of functions apart from its important role in protein syn-
thesis. For instance, so called non-coding RNAs were
found to play a vital role in gene regulatory processes,
which is impressively documented by the discovery of
riboswitch sequences in the 50- and 30-untranslated
regions of mRNA partitions. These RNAs, comprising
an aptamer and expression platform, respond to cellular
concentration levels of metabolites by altering their con-
formations, leading to either up- or down-regulation of a
*To whom correspondence should be addressed. Tel: +43 512 507 5208; Fax: +43 512 507 2892; Email: christoph.kreutz@uibk.ac.at
ß The Author(s) 2011. Published by Oxford University Press.
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/
by-nc/2.5), which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.

Nucleic Acids Research, 2011, Vol. 39, No. 10 4341
monitored by observing the imino proton region of the 1H
NMR spectrum. The refolding kinetics was found to be
remarkably slow with apparent exchange rates (kex) as low
as 0.01 sꢀ1 (11–14).
solvents were extensively dried using freshly activated
˚
molecular sieves (4 A).
50-O-(4,40)-Dimethoxytrityl-20-O-[13C]-methyluridine (2)
Wenter et al. (15) also applied 1H-detected 15N exchange
NMR spectroscopy to characterize refolding kinetics of a
34 nt RNA. This sequence had been designed to undergo
exchange on a time scale accessible to characterization by
longitudinal exchange spectroscopy (<5 s). A major
drawback of the presented approach is the dependence on
the exchangeable imino protons, thus water exchange rates
have to be determined and considered in the analysis of the
exchange process (15).
A
suspension of 50-O-DMT-anhydrouridine
1 (1 g,
1.9 mmol) was suspended in anhydrous dimethyl-
formamide (30 ml). Then, freshly prepared magnesium
[13C]-methoxide (6 eq., 977 mg, 11 mmol) was added.
The mixture was heated to 100ꢁC for 2 h. After evapor-
ation of the solvent, the residue was suspended in
ethylacetate and washed twice with saturated sodium bi-
carbonate solution. The organic phase was then washed
with water, dried over sodium sulfate and finally the
solvent was evaporated. After drying under high vacuum
compound 2 was used without further purification in the
next step. Yield: 987 mg of 2 as white foam (93%). TLC
(CH2Cl2/CH3OH, 95/5): Rf = 0.7; 1H NMR(500 MHz,
CDCl3): d 2.60 (d, J = 9 Hz, 1H, HO-C(30)); 3.64
Here, we report a straightforward labeling strategy
using 20-O-13CH3 modified uridine as a very sensitive
reporter group to study various RNA conformations
and the refolding kinetics thereof. Using 13C longitudinal
exchange NMR spectroscopy we were able to characterize
the refolding of the aforementioned bistable RNAs
(8,12,13,16,17). Furthermore, the dynamic behavior of
an RNA sequence coexisting in three folded states, i.e.
two hairpin-folds and a pseudoknot conformation, was
investigated by Carr–Purcell–Meiboom–Gill (CPMG) re-
laxation dispersion NMR spectroscopy (18–21). To date,
only a limited number of CPMG dispersion studies on
RNA exist, the reason being that the experiments per-
formed in uniformly 13C-enriched RNA suffer from arti-
facts introduced by scalar couplings between adjacent
carbons (21–24). This problem was recently circumvented
13
(d, JCH = 145 Hz, 3H, H3 CꢀO(20)); 3.59 (m, 2H,
H2-C(50)); 3.81 (m, 7H, H-C(20) and 2 OCH3); 4.00 (m,
1H, H-C(40)); 4.51 (m, 1H, H-C(30)); 5.28 (d, J = 8.5 Hz,
1H, H-C(5)); 5.98 (d, J = 1 Hz, 1H, H-C(10)); 6.86 (m, 4H,
H-C(ar)); 7.27–7.41 (m, 9H, H-C(ar)); 8.06 (d, J = 8.5 Hz,
1H, H-C(6)); 8.50 (s, br, 1H, NH) ppm; 13C NMR
(125 MHz, CDCl3): d 55.1 (OCH3); 58.6 (20-O-13CH3);
68.3 (C(30)); 83.5 (C(40)); 84.0 (C(20)); 87.1 (C(10)); 102.2
(C(5)); 113.8 (C(ar)); 128.4, 128.6, 130.4 (C(ar)); 140.0
(C(6)) ppm.
13
13
by Johnson et al. (25) introducing a C20-, C40-labeling
50-O-(4,40)-Dimethoxytrityl-20-O-[13C]-methyluridine
30-(2-cyanoethyl)-N,N-diisopropyl-phosphoramidite (3)
methodology eliminating perturbing
1JCC-effects.
Alternatively, Al-Hashimi and co-workers (26) introduced
an off-resonance 13C R1r experiment to study nucleic
acid dynamics with exchange life times of 10 ms. With
the introduction of the 20-O-13CH3 modified uridine
moiety we create a magnetically isolated spin system
where CPMG type experiments can be performed in a
straightforward way (27). In addition, the approach
allows the site-specific modification of the RNA of
interest.
Compound 2 (530 mg, 0.94 mmol) was dissolved in a
mixture of anhydrous methylene chloride (6 ml) and
N-ethyldimethylamine (10 eq., 1020 ml, 9.4 mmol). After
5 min stirring at room temperature, 2-cyanoethyl-N,
N-diisopropyl-chlorophosphoramidite (1.5 eq., 335 mg,
1.4 mmol) was added dropwise. Stirring was continued
for 2.5 h at room temperature. Then a few drops of
methanol were added and stirring was again continued
for 30 min. The reaction mixture was diluted with methy-
lene chloride and then washed with saturated sodium
bicarbonate solution. The organic phase was dried over
sodium sulfate, evaporated and the residue dried under
high vacuum. The crude product was purified by column
chromatography (CC) on silica (hexanes/ethylacetate
50/50 to 40/60 plus 1% NEt3). Yield: 504 mg of 3
(mixture of diastereoisomeres) as white foam (71%). TLC
(hexanes/ethylacetate 30/70): Rf = 0.7, 0.5; 1H NMR
MATERIALS AND METHODS
Synthesis of the 20-O-13CH3 modified uridine
phosphoramidite (3)
General. NMR spectra were acquired on a Varian
500 MHz Unity Plus instrument. Chemical Shifts are
reported relative to TMS and referenced to residual
proton solvent signal: CDCl3 (7.26 ppm) for 1H NMR
spectra and CDCl3 (77.0 ppm) for 13C spectra. 31P shifts
are reported relative to external phosphoric acid (85%).
1H assignments are based on COSY experiments. 13C
shifts are assigned from a gradient selected phase sensitive
HSQC experiment. Silica 60F-254 plates were used for
thin layer chromatography (TLC). For flash column chro-
matography (FC) silica gel 60 (230–400 mesh) was used.
All reactions were conducted under argon atmosphere.
Reagents and solvents were purchased from Sigma-
Aldrich and used without further purification. Organic
(500 MHz,
CDCl3):
d
1.03ꢀ1.29
(m,
24H,
((CH3)2CH)2N); 2.44, 2.66 (2 m, 4 H, CH2CN); 3.45ꢀ3.61
(m,
16 H,
((CH3)2CH)2N,
POCH2,
H2ꢀC(50),
13
H3 CꢀO(20)); 3.80 (2s, 12H, OCH3); 3.91 (m, 2H,
H-C(20)); 3.96 (m POCH2); 4.21, 4.63 (2m, 2H, HꢀC(40));
4.52, 4.66 (2m, 2H, H-C(30)); 5.21, 5.24 (2d, 2H, J = 8.0 Hz,
HꢀC(5)); 5.99, 6.01 (d, J = 1 Hz, 2H, HꢀC(10)); 6.86 (m,
8H, HꢀC(ar)); 7.25ꢀ7.43 (m, 18H, HꢀC(ar)); 7.99, 8.08 (d,
2H, J = 8.0 Hz, HꢀC(6)) ppm; 31P NMR (121 MHz,
CDCl3): d 151.47, 150.98 ppm; ESI-MS (m/z): [M+H]+
calcd for [13C1]-C40H49N4O9P, 761.80; found 761.87.

4342 Nucleic Acids Research, 2011, Vol. 39, No. 10
Solid phase synthesis and deprotection of RNA sequences
1H NMR spectra of H2O-samples were acquired using a
double-pulsed field gradient spin-echo (DPFGSE) pulse
sequence (29). Assignments of the imino proton reson-
ances for bistable sequences 4 and 5 were reported else-
where (12,30). Assignments of imino proton resonances
for sequences 6, 6a and 6b were achieved by jump-
and-return homonuclear 1H-NOESY experiments (31).
The 13C labeled uridine phosphoramidite was used in
combination with standard 20-O-TOM protected
building blocks (GlenResearch, VA, Sterling) to synthe-
size RNA sequences 4, 5 and 6 and the corresponding
reference sequences. Custom primer supports PS 200
(GE Healthcare) with an average loading of 80 mmol gꢀ1
1
Gradient selected phase sensitive H, 13C-HSQC spectra
were used. The sequences were synthesized on
a
were recorded using a standard pulse sequence (with
optional WATERGATE water suppression for H2O
samples).
Pharmacia Gene Assembler Plus using RNA standard
methods. Amidite (0.1 M) and activator (5-benzylthio-
1H-tetrazole, 0.25 M) solutions were dried over freshly
activated molecular sieves overnight.
Longitudinal exchange experiments are based on pulse
1
sequences previously published for 15N, yielding 13C, H
The removal of protecting groups and the cleavage from
solid support was achieved by treatment with aqueous
methylamine (40%, 650 ml) and ethanolic methylamine
(8 M, 650 ml) at room temperature for 8 h. After evap-
oration the 20-O-protecting groups were removed by
adding 1 M tetrabutylammonium fluoride (TBAF) in
THF (1200 ml). After 14 h at 310 K the reaction was
quenched by the addition of 1 M triethylammonium
acetate (TEAA, pH 7.0, 1200 ml). The volume was
reduced to ꢂ1 ml and then applied on a HiPrep 26/10
desalting column (GE Healthcare). The crude RNAs
were eluted with water, evaporated to dryness and
dissolved in 1 ml water.
2D correlation maps with amplitude modulation of cor-
relation- and exchange-peaks determined by longitudinal
exchange rate constants and the kinetics of interconver-
sion (16). For a ꢂ1 mM sample of sequence 4, arrays of
spectra were recorded at 18.8 T at 298, 300, 303, 306 and
310 K with mixing periods of 5, 100, 200, 300, 400, 600,
800 and 1000 ms (the last data point with 1000 ms was
omitted at 298 K). The size of the data matrices for each
spectrum was 819*64 complex data points, the number of
scans was 48 and the interscan delay was 1.2 s (1 s at
298 K) to yield a total measuring time of 23 h at each
temperature (17 h at 298 K). Arrays of longitudinal
exchange experiments for sequence 5 were recorded at
18.8 T for three temperatures (312, 314 and 316 K) and
analyzed at 314/316 K. Mixing times were 5, 100, 200,
300, 400, 400, 500, 600, 700, 800, 900 and 1000 ms, respect-
ively. Data matrices were 1024*18 (the spectral width in
the carbon dimension was set to 160 Hz), the number of
transients was set to 48 and the interscan delay was 2.6 s,
yielding a total experimental time of 20.5 h.
Analysis and purification of RNA sequences
The quality of the crude RNAs was checked via anion
exchange chromatography on a Dionex DNAPac PA-100
column (4 ꢃ 250 mm) using standard eluents and at
elevated temperature (80ꢁC) (28). Purification of the
RNA sequence of interest was achieved by applying the
crude RNA on a semi-preparative Dionex DNAPac
PA-100 column (9 ꢃ 250 mm) using the standard protocol
(28). The fractions containing the desired RNA were
pooled and loaded on a C18 SepPak catridge (Waters) to
remove HPLC buffer salts. The RNA triethylammonium
salt form was then eluted from the C18 column with water/
acetonitrile (1/1, v/v) and lyophilized. To obtain the RNA
sodium salt form, the RNAs were treated with a sodium
chloride solution (250 mM, 2 ꢃ 10 ml) and water
(3 ꢃ 10 ml) in a Vivaspin 20 ultra-centrifugation unit with
a molecular weight cutoff of 3000 (Satorius Stedim). The
integrity of the RNAs was further checked by mass spec-
troscopy on a Finnigan LCQ Advantage MAX ion trap
instrumentation connected to an Amersham Ettan micro
LC (GE Healthcare).
13C CPMG relaxation dispersion experiments on RNA
sequence 6 were recorded at 11.7 and 18.8 T using the
pulse sequence published by Skrynnikov et al. (27) at
300 K with CPMG field strengths of 33.3, 33.3, 66.7,
100, 133.3, 166.7, 200, 233.3, 266.7, 266.7, 300, 333.3
400, 466.7, 600, 600, 700 and 800 Hz. The relaxation
delay was 60 ms. 1024*18 (1501*17) complex data points
were recorded at 18.8 (11.7) T with the spectral widths in
the proton and carbon dimensions set to 5000 and
150.9 Hz (3000 and 94 Hz), respectively. The number of
transients was set to 100 (18.8 T) and 92 (11.7 T) and the
interscan delay was 2.9 s, yielding a total experimental
time of 55 h (18.8 T) and 45 h (11.7 T).
Data analysis
Spectral processing and peak integration were performed
using NMRPipe and the NMRDraw software package
(32). All subsequent steps were performed using in-house
written software written in Matlab (The MathWorks,
www.mathworks.com).
NMR spectroscopy
The NMR samples were prepared by lyophilization of the
RNA sodium salt form. The RNAs were dissolved
together with 50 mM sodium arsenate buffer, pH 6.5 in
H2O/D2O 9/1 (sequence 4, S4a, S4b, 6a and 6b) or pure
D2O (sequences 5 and 6). The NMR samples were then
heated to 353 K and snap-cooled on ice.
NMR data were acquired on a Varian ‘Unity Plus’ in-
strument operating at 11.7 T or on Varian ‘Inova’ spec-
trometers operating at 11.7 and 18.8 T at the temperatures
indicated below.
Longitudinal exchange experiments. A numerical fitting
procedure was employed for the analysis of data
obtained of sequences 4 and 5 with the minimization
2
qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi
parameter being ꢀ2 ¼ hðIexp ꢀ IcalcÞ i. Errors in the
extracted rate constants were determined by Monte
Carlo analysis where peak intensities were randomly

Nucleic Acids Research, 2011, Vol. 39, No. 10 4343
spectroscopy. This is particularly important for studying
the kinetics of interconversions of the RNAs investigated
here as their conformers are expected to interconvert
rather slowly; for instance, the exchange between two
distinct states of a 34 nt hairpin structure has been
shown to occur with refolding rates ranging from
0.00003 to 0.013 sꢀ1 at 288 and 313 K (11).
modulated according to noise levels in the 2D correlation
maps. Since we approximated peak volumes by summing
over 5 ꢃ 5 (t1 ꢃ t2) data points we assumed that initial
peak intensities are directly related to the populations of
the conformers (33). This approach is justified by compari-
son to a fitting protocol where initial intensities were
fit along with the other parameters. We opted to use the
simplified procedure with fixed initial intensities in order
to avoid numerical instabilities associated with the larger
number of fit parameters. In order to qualitatively account
for cross-correlated relaxation of the correlation peaks,
the numerical fit was extended to a 4 ꢃ 4 matrix
comprising an additional ‘effective’ magnetization mode
for each species that interconvert with the original mag-
netization mode due to an effective cross-correlated relax-
ation rate (Supplementary Table S3). This approach was
sufficient to ensure the decays of the correlation peaks to
be fit correctly, with results virtually unaltered with
respect to a numerical approach that disregards cross-
correlated relaxation.
We decided to attach the 13C-methyl group to the
20-hydroxyl group of the uridine ribose. This labeling
scheme was chosen for several reasons: (i) the building
block is easily accessible by means of chemical synthesis
and (ii) it yields high signal-to-noise ratios in NMR
spectra due to the aforementioned favorable relaxation
properties. Most importantly, (iii) the label represents a
minimally invasive perturbation of the RNA sequence
under investigation. The 20-methoxy modification is a con-
stituent of naturally occurring RNAs, for instance of
transfer ribonucleic acids, and has little influence on the
native conformation of a given RNA. The modification
does not interfere with the C30-endo sugar conformation
favored by RNA and has only a small stabilizing effect
resulting in a slightly higher melting temperature (ꢂ1–2 K
per modification) (36). However, we are aware that the
major drawback of the 20-O-13CH3-label is the deletion
of the 20-hydroxyl group, which is important, for
instance, in the formation of a hydration spine in the
minor groove of double helical RNA structures.
Furthermore, the OH-group can be part of non-canonical
RNA structural elements, e.g. tetraloops (37–39). The
problem can be circumvented by a careful choice of the
labeling position, as discussed below.
CPMG relaxation dispersion. To assess the time-scale of
the dynamics of multi-stable RNA 6, we employed general
numerical two-state fits as well as approximations for very
slow and very fast exchange on the chemical shift
time-scale (34,35). Initial conditions were obtained by an
extensive grid search. The goodness of fit was assessed as
qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi
ꢀ2 ¼ hðRe2xp ꢀ R2calcÞ =ꢁ2i, where s is the standard devi-
2
ation computed from duplicate experiments. In addition,
the reduced ꢀ2 value was computed for each model. The
significance of the respective models was tested with
respect to fitting a flat line using F statistics. For every
model, a Monte Carlo analysis was performed where
errors were estimated on the basis of the duplicate experi-
ments. In addition, a restricted three-state fit was per-
formed with populations and chemical shift differences
Synthesis of 20-O-13CH3-labeled uridine phosphoramidite
building block
The synthesis of the 13C labeled uridine building block
started from 50-O-DMT-20-deoxyanhydrouridine 1,
which was prepared according to published procedures
(Scheme 1) (40). Compound 1 was quantitatively con-
verted into the 20-O-13CH3-labeled intermediate 2 by treat-
ment with Mg(O13CH3)2, which was freshly prepared
from magnesium turnings and 13C labeled methanol in
analogy to earlier works using unlabeled methanol (41).
Subsequent phosphitylation with (2-cyanoethyl)-N,
1
constrained at values obtained from 13C, H correlation
maps and assuming only indirect interconversion between
conformers 600 and 6000 (via 60) (for details see
Supplementary Table S5).
N-diisopropylchlorophosphoramidite
provided
the
Arrhenius analysis. The temperature dependence of the
equilibrium constant and the rate constants for sequence
4 obtained from longitudinal exchange data was analyzed
by linear regression of ln (K) versus 1/T and ln (kij) versus
1/T, respectively. The equilibrium and activation param-
eters were computed in each Monte Carlo run yielding the
corresponding mean values and standard deviations.
desired 20-O-13CH3-labeled uridine building block 3 with
RESULTS AND DISCUSSION
Choice of 13C label
Scheme 1. Synthesis of the 20-O-13CH3-uridine phosphoramidite 3;
(a) 6 eq. 13C-magnesium methoxide [Mg(O13CH3)2, freshly prepared from
magnesium turnings and 13CH3OH] in anhydrous dimethylformamide,
100ꢁC, 2 h, 93%; (b) 1.5 eq. (2-cyanoethyl)-N,N-diisopropylchloro-
phosphoramidite, 10 eq. N-ethyldimethylamine in anhydrous dichloro-
methane, room temperature, 2.5 h, 71%.
One important reason for the choice of this particular spin
label is the comparatively slow relaxation of methyl 13C
magnetization, which increases the time-window where
slow refolding processes are accessible to observation
and analysis by longitudinal exchange NMR

4344 Nucleic Acids Research, 2011, Vol. 39, No. 10
66% overall yield in two steps with one chromatographic
purification step. The 13C building block 3 was then used
in RNA solid-phase synthesis and was incorporated with
>98% coupling efficiency. The molecular weights of
the 13C-labeled RNAs were confirmed by LC-ESI mass
spectroscopy (Supplementary Table S1).
Supplementary Figure S2). Quantification of the equilib-
rium population distribution was obtained from 2D
peak integration to yield relative populations of folds 50
and 500 of ꢂ7/3 (Figure 1F). Of note, the 13C uridine
label resides in a single stranded region in both conform-
ations. This leads to a less pronounced chemical shift
difference in both dimensions as compared to sequence
4
(Áo1H4 4 = 0.305 ppm, Áo1H5 5 = 0.17 ppm and
0
00
0 00
Labeling strategy, studied RNA sequences and NMR
spectroscopic characterization of the individual folds
Áo13C4 4 = 1.33 ppm, Áo C5 5 = 0.31 ppm). However,
the chemical shift differences between the two states in
the carbon and proton dimensions are still sufficient to
13
0
00
0 00
0
One of the key steps is the correct0 choice of the labeling
position. The 20-O-13CH3-modified uridine was placed in
such a manner that the label is residing in a solvent
accessible single stranded position in one of the folds,
whereas in the competing conformation the 13C-methyl
modified uridine is part of a double helical A-form
hairpin stem. Thereby, the magnetic environment is
expected to be sufficiently different for the chemical
shifts of the competing states to be resolved in an NMR
spectrum. The labeling strategy was chosen in line with
earlier observations (28).
The realization of the concept was demonstrated by
introducing the 13C-uridine 3 into the 20 nt bistable
RNA sequence 4 at position 18 (Figure 1A) (12). We
observed two distinct signal patterns in both the imino
1
1
proton regions of the H-NMR spectrum and in the H,
13C-HSQC spectrum of sequence 4 originating from the
two competing folds, 40 and 400, indicating that the two
species refold slowly on the NMR chemical shift
time-scale (Figure 1B and C). This nicely demonstrates
the sensitivity of the isotope label to its magnetic environ-
ment. The 13C, 1H resonances were assigned to their
respective conformations by the aid of truncated reference
sequences (sequences S4a and S4b, Supplementary
Figure S1). Quantification of the equilibrium fold distri-
bution was checked on one hand by the comparative
imino proton method [for a detailed description of the
comparative imino proton method see reference (8)] and
1
on the other hand by integration of the H, 13C-HSQC
peaks. Both methods reported the two states to be equally
populated, in line with earlier findings (8,12). Importantly,
the 13C-modification also proved to be minimally invasive
as no population differences compared to the wild-type
sequence were found. This was further supported by
UV-melting experiments, which show that the 20-O-
methoxy modification change the melting transition by
only 1–2 K (Supplementary Figure S3) (8).
In order to further explore the applicability of this novel
13C RNA labeling methodology we modified the larger
bistable RNA 5 comprising 32 nt with a 20-O-13CH3-
uridine building block at position 26 (Figure 1D). For
this medium sized RNA sequence 5 severely overlapping
signals in the region between 12.5 and 13.5 ppm of the
1H-NMR spectrum were encountered (Figure 1E). So
far, only partial assignment of the NH resonances has
been achieved by site-specific 15N labeling (30).
This case of resonance overlap demonstrates the poten-
tial of our site-specific 13C labeling strategy. Again, two
well separated 1H, 13C-HSQC resonances are detected
and assigned to the individual folds by comparison with
Figure 1. Realization of the presented approach as exemplified on two
bistable RNAs (4 and 5). (A) Bistable 20 nt RNA 4 with the two
competing folds 40 and 400. The red U denotes the 20-O-13CH3-uridine
label. (B) Detection of the folding equilibrium of conformation 40 and
400 by analysis of the imino proton region of the 1H NMR spectrum.
1
(C) H, 13C-HSQC of RNA sequence 4. The two folding states give rise
to two well-resolved peaks in the HSQC spectrum. Assignment was
achieved by means of truncated reference sequences (Supplementary
Information). (D) Bistable 32 nt RNA 5 with the two competing folds
50 and 500. The red U denotes the 20-O-13CH3-uridine label. (E) Severe
resonance overlap is found in the imino proton region of the 1H NMR
spectrum. (F) 1H, 13C-HSQC spectrum of RNA sequence 5. The two
conformations are nicely resolved in the HSQC spectrum. Fold assign-
ment was achieved using a truncated reference sequence (S5a, see
Supplementary Information). Conditions: 0.8–1.0 mM RNA, 50 mM
sodium phosphate, pH 6.5, H2O/D2O 9/1, 298 K.
a
truncated reference sequence (sequence S5a,

Nucleic Acids Research, 2011, Vol. 39, No. 10 4345
distinguish the two states. This observation further
strengthens the impact of the presented methodology, as
slight deviations from the suggested labeling strategy
(single-stranded versus double-stranded) seem to be
tolerated. The ratio of peak integrals of folds 50 and 500
was 7/3, which is identical to the one given in reference
(30), proving that the label did not affect the thermo-
dynamic equilibrium of the conformations. Differences
in melting temperatures between wild-type and modified
samples determined from UV-melting experiments were
very small with ÁTm (20-O-CH3 versus wild-type) values
between 0.5 and 1 K (T1m (5) = 55.8ꢁC, T2m (5) = 85.8ꢁC
versus T1m (5WT) = 55.0ꢁC, Tm2 (5WT) = 85.3ꢁC and Tm
(S5a) = 70.6ꢁC
versus
Tm
(S5aWT) = 69.8ꢁC,
Supplementary Figure S3) (8). These findings demonstrate
the minimally invasive nature of our modification.
As a final example RNA sequence 6 was chosen (18,19).
The oligonucleotide 6 coexists in three competing folds, a
pseudoknotted conformation 60, a 50-hairpin fold 600 and a
30-hairpin fold 6000 (Figure 2A). The presence of multiple
folding states is not directly observable by the imino
proton resonances, as the NH resonances originating
from all three conformations (É-knot sequence 60,
50-hairpin 600 and 30-hairpin 6000) are strongly overlapping
(Figure 2B).
The coexistence of the three folding states was revealed
1
by the presence of three observable resonances in the H,
13C correlation map of RNA 6 (Figure 2C). By incorpor-
ation of the 20-O-13C-methyl uridine label into sequence 6
at position 16 and into the respective reference sequences
1
(6a and 6b), it was possible to assign the methyl H/13C
resonances. Peak integration of the 1H, 13C-HSQC correl-
ation maps yields an estimate of the individual popula-
tions, namely a ratio of ꢂ4/1/2 of pseudoknot 60,
50-hairpin 600 and 30-hairpin 6000 at 298 K. The population
distribution was found to be strongly temperature depend-
ent according to peak volumes obtained at various tem-
peratures (278, 283, 298 and 303 K, Figure 3B). While the
pseudoknotted conformation 60 was favored at lower tem-
peratures, the hairpin folds 600 and 6000 were increasingly
populated at higher temperatures (Figure 3C). This strong
temperature dependence is indicative of enthalpy–entropy
compensation determining the equilibrium distributions of
the three species. Whereas the pseudoknotted conform-
ation is enthalpically favored (8 bp), it is entropically
destabilized with respect to the hairpin structures that
have long unstructured regions.
Figure 2. (A) Multi-stable RNA sequence 6 is able to fold into a
pseudoknotted conformation 60 and two hairpin structures 600 and
6
000. Truncated reference sequences 6a and 6b mimic fold 600 and 6000
,
respectively. The red U denotes the 20-O-13CH3-uridine label. (B) Imino
proton region of the 1H NMR spectrum of sequence 6 and references
6a and 6b. The detection of multiple conformations of RNA 6 is
hampered due to NH resonance overlap originating from the individual
folds. Asterisks denote unassigned peaks most likely originating from
fold 600 and 6000. (C) 1H, 13C-HSQC of RNA sequence 6. The three
possible folding states can be easily detected and assigned in the
HSQC spectrum. Fold assignment was achieved with truncated refer-
ence sequences 6a and 6b. Here, an overlay of the HSQC spectra of 6a
and 6b is shown. Conditions: 0.75 mM RNA, 2.5 mM MgCl2, 50 mM
sodium phosphate, pH 6.5, H2O/D2O 9/1, 298 K.
of a few seconds can be studied by longitudinal
exchange experiments, whereas even slower rearrange-
ments can be studied by real-time NMR methods (14,43).
First, we have determined the kinetics of intercon-
version between the two magnetic environments of the
uridine 20-O-13CH3 label in the RNA sequence 4 by 13C
longitudinal exchange experiments. This process was
previously studied via a fold-caging-real-time NMR
approach (12), yielding surprisingly slow refolding
Refolding kinetics and conformational dynamics studied
by longitudinal-exchange and CPMG NMR experiments
One of our main goals was to determine the refolding
kinetics of our model systems at unperturbed equilibrium.
RNA sequences 4, 5 and 6 are expected to interconvert
between their conformational substates on time scales
ranging from milliseconds to a few seconds. There are
several NMR-based methods that are suited to character-
ize processes occurring at various time scales (42).
Motions in the millisecond to microsecond time regime
can be characterized using CPMG relaxation dispersion
NMR methods (20). Dynamics occurring on the order
0
00
00
0
kinetics [kex = k4 !4 +k4 !4 ranged between 0.01
(at 283 K) and 0.2 sꢀ1 (298 K)]. These results were used
to benchmark the new labeling scheme and its influence
on the kinetic properties of this RNA sequence (12).

4346 Nucleic Acids Research, 2011, Vol. 39, No. 10
Figure 3. Temperature dependence of folding state populations of RNA
sequence 6. (A) Representation of the three possible folding states. The
red U denotes the 20-O-13CH3-uridine label. (B) 1H, 13C-HSQC spectra of
RNA sequence 6 at various temperatures ranging from 278 to 303 K. (C)
Folding state populations (in %) versus temperature. The populations
were determined as the mean value from three independent HSQC meas-
urements at the indicated temperature (error bars from standard devi-
ation). Conditions: 0.75 mM RNA, 2.5 mM MgCl2, 50 mM sodium
phosphate, pH 6.5, H2O/D2O 9/1, 298 K.
Figure 4. Kinetics of sequence 4 analyzed by 13C longitudinal exchange
spectroscopy. (A) Interconversion between conformations 40 and 400. The
uridine nucleotide that serves as a sensor is highlighted in red. (B) Signal
intensities as a function of mixing time are shown for a set of temperatures.
The left panel shows normalized intensities of the correlation peak of
conformation 40 and the exchange peak corresponding to transition
40 ! 400. The right panel depicts the intensities of the correlation peak of
conformation 400 and the exchange peak corresponding to transition
400 ! 40. Experiments performed at different temperatures are color-coded
(298 K: blue, 300 K: magenta, 303 K: red, 306 K: orange, 310 K: yellow).
1
We recorded a 13C, H resolved longitudinal exchange
experiment which yields a series of 13C, H correlation
1
Table 1. Microscopic rate constants and longitudinal relaxation rates
along with errors obtained by longitudinal exchange experiments con-
ducted on sequence 4 at each temperature analyzed
maps with correlation- and exchange-peaks that have
been amplitude modulated during a mixing time according
to their longitudinal relaxation rates and their kinetics
of interconversion. Decays and buildups at the different
temperatures are shown in Figure 4. Assuming a two-state
process we determined the refolding kinetics in the
temperature range between 298 and 310 K yielding
Temperature (K) k4 !4 (sꢀ1
)
(sꢀ1
)
R1(40) (sꢀ1
)
R1 (400) (sꢀ1
)
00!40
k4
0
00
298
300
303
306
310
0.051 0.002 0.039 0.002 2.13 0.03
0.065 0.003 0.048 0.003 2.26 0.03
0.102 0.007 0.057 0.004 2.46 0.06
0.191 0.006 0.102 0.005 2.59 0.14
0.419 0.010 0.155 0.004 2.30 0.18
1.96 0.04
2.12 0.03
2.20 0.06
2.42 0.16
2.43 0.23
0
00
microscopic exchange rate constants k4 !4 between
0.05 and 0.42 sꢀ1 and k4 !4 between 0.04 and 0.16 sꢀ1
with errors being on the order of 5–10% (Table 1). This
,
00
0
Effective longitudinal relaxation rates are obtained by accounting for
cross-correlated relaxation (‘Materials and Methods’ section and
Supplementary Information).
corresponds to an exchange rate conꢀst1ant kex
0
00
00
0
(kex = k4 !4 +k4 !4 ) between 0.09 and 0.58 s and the
0
00
population distribution of p4 /p4 changing from 0.57/0.43
to 0.73/0.27, which agrees well with the values obtained
from integration of peak volumes (0.50/0.50 to 0.65/0.35
in the corresponding temperature range).
15.0
EA
= 21.6 0.88 kcal molꢀ1/A4#-4 = 10
were
00
4#-400
obtained. The activation energy was found to amount to
55% (42%) of the (base pairing and stacking) enthalpy as
computed by m-fold for fold 40 (400) (44). This makes a
dissociative mechanism, which would imply a fully
unfolded state, rather unlikely, but favors an associative
mechanism with base pairs of both conformational states
formed and being disrupted at the same time during the
transition state (13).
Analysis of the temperature dependence of the equilibrium
constant obtained from the microscopic exchange rates yielded
00
0
=
a
refolding enthalpy and entropy of ÁH4 -4
00
11.2 1.1 kcal molꢀ1 and ÁS4 -4 = 38.1 3.7 cal molꢀ1 Kꢀ1
.
0
The temperature dependence of the equilibrium constant and
the microscopic rate constants are shown in Figure 5.
Thus, the results from the analysis of longitudinal
exchange experiments are in line with those direcꢀtl1y
The bistable RNA 4 was studied before in the tempera-
ture range between 283 and 298 K (12). The rate constants
found at 298 K were compared to those determined previ-
ously at the same temperature and found to bꢀe1in good
00
0
obtained from peak integration (ÁH4 -4 =9.9 kcal mol
,
ÁS4 -4 = 33.1 cal molꢀ1 Kꢀ1). Assuming an Arrhenius
temperature dependence of the rate constants, activa-
tion energies and approximated frequency factors
00
0
agreement (k4 !4 = 0.051 sꢀ1, k4 !4 = 0.039 s iꢀn1 our
0
00
00
0
EA
= 32.8 0.68 kcal molꢀ1/A4#-4 = 10
and
23.1
work versus k4 !4 = 0.091 sꢀ1, k4 !4 = 0.076 s
in
0
4#-40
0
00
00
0

Nucleic Acids Research, 2011, Vol. 39, No. 10 4347
reference (12), see also Supplementary Table S4). The tem-
perature dependence of the equilibrium constant is,
however, more pronounced in our study, indicating
larger enthalpy and entropy differences between the two
conformers than previously reported (12). We find that
fold 400 (which contains the label within the base-paired
region) is enthalpically destabilized with respect to 40.
Over the observed temperature range, fold 400 is,
however, thermodynamically more stable than fold 40,
indicating that fold 400 is entropically favored over 40.
The difference is reflected in the activation parameters of
the backward reaction 400 ! 40, indicating that the
enthalpic portion of the barrier (related to the activation
energy) is smaller than for the forward reaction, whereas
frequency factors indicate a larger entropic barrier. We
believe that the discrepancy is due to the different tem-
perature range and the lower pH employed in our study,
as the effect of the uridine label is expected to only slightly
stabilize secondary structures (see UV melting analysis in
Supplementary Information).
We subsequently applied the longitudinal exchange
approach to the thermodynamically more stable 32 nt
RNA stem loop structure 5 that coexists in two distinct
conformations 50 and 500. In order to ensure that the
reaction was fast enough to be observable by this
approach, experiments were conducted at elevated tem-
peratures. For this RNA it was possible to obtain mean-
ingful results from experiments conducted at 314 and
316 K. Respective correlation- and exchange-peak
intensities as a function of mixing time are shown in
Figure 6. The rate of interconversion at this temperature
is in the same range as for sequence 4 at room tempera-
ture, i.e. k5 !5 = 0.348 0.063 sꢀ1 (0.323 0.056 sꢀ1
)
at
0
00
and k5 !5 = 0.206 0.024 sꢀ1 (0.262 0.024 sꢀ1
)
00
0
314 K (316 K). The complete set of kinetic parameters is
given in Table 2. Due to the more pronounced skew in
populations (approximately 7/3) as compared to RNA
sequence 4, the exchange peak derived from the less
Figure 6. 13C longitudinal exchange experiments conducted on
sequence 5. (A) Interconversion between conformations 50 and 500
with the uridine label highlighted in red. (B) Left panel: intensities of
the corrrelation peak corresponding to conformation 50 and exchange
peak corresponding to transition 50 ! 500 as a function of mixing time.
Right panel: correlation peak pertinent to fold 500 and exchange peak
for the transition 500 ! 50. Results at 314 (316) K are depicted as circles
(diamonds) and fits at 314 (316) K are shown as solid/dashed lines.
Error bars were obtained on the basis of spectral noise in a Monte
Carlo analysis.
Table 2. Microscopic rate constants and longitudinal relaxation rates
along with errors obtained by longitudinal exchange experiments con-
ducted on sequence 5 at each 314 and 316 K
Figure 5. Analysis of the refolding reaction of sequence 4 in terms of
kinetic and thermodynamic parameters obtained from longitudinal
exchange experiments on the 20-O-13CH3-uridine label. (A) ln (k4 !4
0
00
)
Temperature (K) k5 !5 (sꢀ1
)
k5
(sꢀ1
)
R1 (50) (sꢀ1
)
R1 (500) (sꢀ1
)
00
0
(solid line) and ln (k4 !4 ) (dashed line) as a function of the inverse
0
00
00!50
0
00
00
0
temperature. (B) ln (K) [computed as ln (k4 !4 /k4 !4 )] as a function
of 1/T. Error bars were obtained on the basis of duplicate data points
by a Monte Carlo analysis, and regression was performed on averaged
314
316
0.348 0.053 0.206 0.024 0.67 0.11
0.323 0.056 0.262 0.024 0.92 0.11
0.88 0.05
0.88 0.05
0
00
00
0
values of k4 !4 , k4 !4 , and K.

4348 Nucleic Acids Research, 2011, Vol. 39, No. 10
populated species displays relatively low intensities in the
longitudinal exchange experiments, which, in conjunction
with a small rate of interconversion, impeded the acquisi-
tion of data of satisfactory signal-to-noise ratio below
314 K. On the other hand, rapid sample degradation
and thermal denaturing effects did not permit extended
exposure of this RNA to higher temperatures. The tem-
perature range investigated for sequence 5 was thus too
narrow to allow reliable determination of thermodynamic
and kinetic information. For even more stable RNA struc-
tures and/or more skewed populations it is advisable to
resort to real-time NMR techniques (14).
Application of longitudinal exchange experiments to
the tristable RNA sequence 6 proved impractical due to
spectral overlap of conformations 600 and 6000 in the proton
dimension. In an attempt to obtain information about the
kinetics of interconversion between the individual con-
formations we conducted 13C CPMG relaxation disper-
sion measurements (21,27). CPMG relaxation dispersion
experiments exploit the phenomenon that resonances
show increased transverse relaxation rates if their
chemical shifts are modulated by some conformational
or chemical process. They permit the characterization of
microsecond-to-millisecond motions and have become
increasingly popular for detecting excited states, which
are not sufficiently populated to be directly observable
in NMR spectra. In the favorable case of exchange
processes occurring in the intermediate regime on the
chemical shift time-scale, the exchange rate constant kex
(which is given by the sum of the forward and backward
rate constants), the population of the individual states pi,
and the chemical shift difference between the states, Áo,
can be obtained, as well as the relaxation rates at infinite
CPMG field strength, R21(i, x), where i stands for the
resonance and x stands for the spectrometer field
strength. In the case of very slow or very fast exchange
processes, not all exchange parameters can be obtained
separately. If kex >> Áo (fast exchange regime), kex can
be accurately determined, along with the product of the
populations and the square of the chemical shift differ-
ences, papbÁo2 and R21(i, x). If exchange is slow on the
chemical shift time-scale (kex << Áo) and the populations
of both individual states are high enough to produce ob-
servable resonances, characteristic relaxation dispersion
profiles are observed that can be analyzed as in
Tollinger et al. (34). In this case, it is not possible to
obtain the ‘backward’ rate constant (or the population
of the excited state), such that only four parameters are
obtained [the ‘forward’ rate constant kab, Áoab, and the
two relaxation rates R12 (i, x)].
Figure 7. Analysis of CPMG relaxation dispersion of RNA sequence 6.
A two-state process that is fast on the chemical shift time-scale was fit
to the data of conformer 60 (A) and 6000 (B) (for details of the fitting
procedure see text and Supplementary Information). Experimental data
points are shown as black circles and the fit is depicted as a red line.
Spectrometer field strengths are indicated and the secondary structures
of the two states are shown as inserts. Both conformers 60 and 6000
exchange with their respective excited state at a rate constant of
ꢂ500 sꢀ1
.
From the magnetic field dependence of the exchange
contribution to transverse relaxation in conjunction with
the goodness of the fit of other models we concluded that
in fact fast two-state exchange was suited to explain
our experimental data (45). According to this model,
conformers 60 (6000) access their excited states at rate con-
stants of 593 133 (460 268) sꢀ1. From the fitted
product function papbÁo2 it follows that, depending
on the relative populations, the chemical shift difference
between the exchanging states are around 0.45 ppm (at an
excited state population of 1%) and 0.8 ppm (at equimolar
populations) for both conformers. The exact results are
given in Table 3.
These results are surprising given the fact that we have
obtained exchange peaks in a 13C-13C resolved 3D ZZ
exchange experiment similar to the experiment conducted
by Sprangers et al. (17) at mixing times of 5, 100 and
150 ms, indicating that species 60 and 6000 interchange on
the order of 3 sꢀ1 (Supplementary Figure S6). A process
We recorded experiments at 300 K at 125 and 200 MHz
carbon frequency. Small but statistically significant disper-
sion profiles could be found for conformers 60 and 6000 (the
least populated conformer 600 could not be analyzed due to
low signal to noise ratio, Figure 7). In order to assess
whether the underlying process is the slow exchange
between the visible conformers, we analyzed the data
using equations for two-state processes on the very slow,
the intermediate and the fast time regimes (34,35). Also
slow exchange in 2 three-state exchange models was
investigated (Supplementary Table S5).

Nucleic Acids Research, 2011, Vol. 39, No. 10 4349
Table 3. Results from CPMG relaxation dispersion experiments performed on sequence 6 at 300 K: two-state models
Two-state models
kex,6*-6** (sꢀ1
)
k6*-6** (sꢀ1
)
P**
Áo (ppm)
P*P** Áo2 (sꢀ2) (125 MHz)
ꢀ2red
Conformer 60
Slow (1)
Slow (2)
Intermediate
Fast
Conformer 6000
Slow (1)
Slow (2)
2.92 0.27
2.53 0.13
0.54 0.05
1.10 0.08
0.13 0.09
0.61
1.21
3.13a
0.34
593 133
579 62
0.20 0.34
0.58 0.32
1098 90
3.79 0.95
2.70 0.50
0.46 0.12
1.07 0.04
0.14 0.10
0.30
0.42
1.82a
0.27
Intermediate
Fast
445 300
460 268
1182 461
Results are derived from 1000 Monte Carlo runs. The two states are tagged by asterisks, where one asterisk refers to the conformer which
corresponds to the visible ground state (60 or 6000), and two asterisks represent the invisible excited state that pertains to it. For both analyzed
resonances, the equation for slow exchange gave two solutions (1) and (2).
aꢀ2redvalues that are larger in the Monte Carlo analysis in comparison to the direct analysis. These discrepancies originate from the fact that the set of
mean values derived from the Monte Carlo analysis does not represent the dispersion curve.
on this time-scale with the given populations and chemical
shift differences would indeed lead to a dispersion profile
of the observed size, albeit, with a different shape. From
this discrepancy we conclude that we are in fact con-
fronted with a mixture of several processes, i.e. the slow
interconversion between conformers 60 and 6000 and their
respective excursions to invisible conformers on the time
scale of several hundred per second. For the parameters in
our system it can be shown by numerical simulation that
the dispersion profile obtained for a mixture of processes
coincides within error with the one that would be obtained
by a purely fast process (Supplementary Figure S7). Thus,
it was not possible by the means of our data to separate
the slow process, i.e. the interconversion between 60 and
6000, from the dominant fast process. On the other hand,
we may safely assume that the resulting parameters of the
fast process have not been significantly perturbed by the
presence of the slower one.
Given the positioning of the label it is tempting to infer
that the observed excursion to an excited state might arise
from the interconversion between different sugar puckers.
However, such sugar pucker dynamics were recently
determined to occur with exchange rates between 20 000
and 40 000 sꢀ1, i.e. about 40 times faster than the process
we observe (21). Given this significant discrepancy, we
tend to believe that the process we observe is not due to
this fast conformational fluctuation but rather results
from a local conformational dynamics in the 9 nt loop
region (this structural feature is intact in both analyzed
conformations). Further experiments using different
label positions and/or types of labels are planned in
order to investigate the nature of this dynamic event.
First, we used the label to assign and quantify fold
populations of multi-stable RNAs. The 13C-modification
proved to be minimally invasive, as no population changes
and only minor melting point changes were observed
compared to the wild-type sequences (8). The refolding
kinetics of RNA 4 could be determined using a 13C longi-
tudinal exchange experiment at temperatures ranging
from 298 to 310 K. The obtained rate constants were in
good agreement with previous findings (12). The subse-
quent Arrhenius analysis yielded an estimation of the tran-
sition state energy between the two competing secondary
structures.
We extended the approach to the more stable RNA
sequence 5 comprising 32 nt. As expected, exchange
peaks were found only at elevated temperature (>313 K).
It was possible to determine the folding kinetics at 314
and 316 K. Not only the velocity of the process but also
the skew in populations poses a limitation to the sensi-
tivity of this approach. For more stable structures or
bistable RNAs with very skewed populations, it is advis-
able to address the refolding kinetics by real-time NMR
methods as previously shown (11).
For the tristable RNA 6 longitudinal exchange experi-
ments were impractical due to poor chemical shift disper-
sion in the proton dimension. As an alternative approach
we used 13C CPMG relaxation dispersion NMR spectros-
copy. Relaxation dispersion data pointed to a process that
is fast on the chemical shift time-scale with kex ꢂ500 sꢀ1
and Áo ꢄ 0.3 ppm for at least two of the species. A slow
process, detected by 13C-13C resolved ZZ-exchange experi-
ments, is obscured by the dispersion profile of the fast
process.
Our 20-O-13CH3-uridine label has proven to be generally
applicable to study conformational heterogeneity and the
refolding kinetics in small to mid-sized RNAs comprising
up to ꢂ35 nt using standard NMR techniques. For even
larger molecules, the signal-to-noise limitation could be
alleviated by using cryogenic probes along with TROSY
variants of the NMR experiments (17,46). The labeling
procedure itself is feasible for RNAs of up to 150 nt
using the recently developed approach that combines
CONCLUSIONS
We used a 20-O-13CH3-uridine label to study RNA refold-
ing kinetics in an unperturbed equilibrium state, thereby
circumventing a major disadvantage of methods that rely
on the observation of exchange labile protons and
enabling the study of RNA refolding phenomena at
elevated temperatures (15).

4350 Nucleic Acids Research, 2011, Vol. 39, No. 10
9. Micura,R. and Hobartner,C. (2003) On secondary structure
rearrangements and equilibria of small RNAs. ChemBioChem, 4,
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10. Schultes,E.A. and Bartel,D.P. (2000) One sequence, two
ribozymes: implications for the emergence of new ribozyme folds.
Science, 289, 448–452.
11. Furtig,B., Wenter,P., Reymond,L., Richter,C., Pitsch,S. and
Schwalbe,H. (2007) conformational dynamics of bistable RNAs
studied by time-resolved NMR spectroscopy. J. Am. Chem. Soc.,
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12. Wenter,P., Furtig,B., Hainard,A., Schwalbe,H. and Pitsch,S.
(2006) A caged uridine for the selective preparation of an RNA
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chemical RNA synthesis and enzymatic ligation methods
(47,48).
Compared to other non-native NMR labeling
approaches for RNA, like incorporation of 5-F-uridine,
we think that our presented approach is as minimally
invasive as the fluorine tag (49–52). Furthermore, we
consider the 20-O-13CH3 labeling protocol more versatile
as triple resonance probes to carry out the 13C-based
experiments are more disseminated in the biomolecular
NMR community than probes suitable for the 19F-based
experiments.
In principle, similar 20-O-13CH3 labeling schemes can be
applied to all 4 nt types conveying a high level of versatil-
ity to the labeling approach. It will also be very interesting
to introduce the 13C-methoxy-residue into RNAs where
this very modification occurs naturally. Subsequent
NMR spectroscopic analysis can be used to gain interest-
ing insight into the structure and function of the methoxy-
modification.
16. Farrow,N.A., Zhang,O., Forman-Kay,J.D. and Kay,L.E. (1995)
Comparison of the backbone dynamics of a folded and an
unfolded SH3 domain existing in equilibrium in aqueous buffer.
Biochemistry, 34, 868–878.
SUPPLEMENTARY DATA
Supplementary Data are available at NAR Online.
17. Sprangers,R., Gribun,A., Hwang,P.M., Houry,W.A. and Kay,L.E.
(2005) Quantitative NMR spectroscopy of supramolecular
complexes: dynamic side pores in ClpP are important for product
release. Proc. Natl Acad. Sci. USA, 102, 16678–16683.
18. Puglisi,J.D., Wyatt,J.R. and Tinoco,J.I. (1990) Conformation of
an RNA pseudoknot. J. Mol. Biol., 214, 437–453.
19. Puglisi,J.D., Wyatt,J.R. and Thomas,L.J. (1995) Methods in
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magnetic resonance methods for quantifying microsecond-to-
millisecond motions in biological macromolecules. Methods
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ACKNOWLEDGEMENTS
C.K. thanks Ronald Micura (Innsbruck) and Bernhard
Krautler (Innsbruck) for continuous support and scientific
discussions. K.K. was a recipient of an APART fellowship
of the Austrian Academy of Sciences at the time when this
study was realized.
FUNDING
21. Johnson,J.E. and Hoogstraten,C.G. (2008) Extensive backbone
dynamics in the GCAA RNA tetraloop analyzed using
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FWF ( V173 to K.K., SFB-17 and P20549 to R.K.).
Funding for open access charge: FWF - Austrian
Science Fund.
Conflict of interest statement. None declared.
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