Structures of small molecules bound to RNA repeat expansions that cause Huntington's disease-like 2 and myotonic dystrophy type 1

Abstract

Trinucleotide repeat expansions fold into long, stable hairpins and cause a variety of incurable RNA gain-of-function diseases such as Huntington’s disease, the myotonic dystrophies, and spinocerebellar ataxias. One approach for treating these diseases is to bind small molecules to the structured RNAs. Both Huntington’s disease-like 2 (HDL2) and myotonic dystrophy type 1 (DM1) are caused by a r(CUG) repeat expansion, or r(CUG)exp. The RNA folds into a hairpin structure with a periodic array of 1×1 nucleotide UU loops (5’CUG/3’GUC; where the underlined nucleotides indicate the Us in the internal loop) that sequester various RNA-binding proteins (RBP) and hence the source of its gain-of-function. Here, we report NMR-refined structures of single 5’CUG/3’GUC motifs in complex with three different small molecules, a di-guandinobenzoate (1), a derivative of 1 where the guanidino groups have been exchanged for imidazole (2), and a quinoline with improved drug-like properties (3). These structures were determined using nuclear magnetic resonance (NMR) spectroscopy and simulated annealing with restrained molecular dynamics (MD). Compounds 1, 2, and 3 formed stacking and hydrogen bonding interactions with the 5’CUG/3’GUC motif. Compound 3 also formed van der Waals interactions with the internal loop. The global structure of each RNA-small molecule complexes retains an A-form conformation, while the internal loops are still dynamic but to a lesser extent compared to the unbound form. These results aid our understanding of ligand-RNA interactions and enable structure-based design of small molecules with improved binding affinity for and biological activity against r(CUG)exp. As the first ever reported structures of RNA r(CUG) repeats bound to ligands, these structures can enable virtual screening campaigns combined with machine learning assisted de novo design.

Full text

NMR structures of small molecules bound to a model of an RNA
CUG repeat expansion
Jonathan L. Chen1,2,†, Amirhossein Taghavi3,†, Alexander J. Frank4, Matthew A. Fountain4,
Shruti Choudhary1, Soma Roy3, Jessica L. Childs-Disney3, Matthew D. Disney*,1,3
1Department of Chemistry, The Scripps Research Institute, 130 Scripps Way, Jupiter, FL 33458,
USA
2Department of Biochemistry and Biophysics, Center for RNA Biology, University of Rochester
Medical Center, Rochester, NY 14642, USA
3Department of Chemistry, The Herbert Wertheim UF Scripps Institute for Biomedical Innovation
& Technology, 130 Scripps Way, Jupiter, FL 33458, USA
4Department of Chemistry and Biochemistry, State University of New York at Fredonia, Fredonia,
NY 14063, USA
Abstract
Trinucleotide repeat expansions fold into long, stable hairpins and cause a variety of incurable
RNA gain-of-function diseases such as Huntington’s disease, the myotonic dystrophies, and
spinocerebellar ataxias. One approach for treating these diseases is to bind small molecules to
the structured RNAs. Both Huntington’s disease-like 2 (HDL2) and myotonic dystrophy type
1 (DM1) are caused by a r(CUG) repeat expansion, or r(CUG)exp. The RNA folds into a
hairpin structure with a periodic array of 1×1 nucleotide UU loops (5’CUG/3’GUC; where the
underlined nucleotides indicate the Us in the internal loop) that sequester various RNA-binding
proteins (RBP) and hence the source of its gain-of-function. Here, we report NMR-refined
structures of single 5’CUG/3’GUC motifs in complex with three different small molecules, a
di-guandinobenzoate (1), a derivative of 1 where the guanidino groups have been exchanged for
imidazole (2), and a quinoline with improved drug-like properties (3). These structures were
determined using nuclear magnetic resonance (NMR) spectroscopy and simulated annealing with
restrained molecular dynamics (MD). Compounds 1, 2, and 3 formed stacking and hydrogen
bonding interactions with the 5’CUG/3’GUC motif. Compound 3 also formed van der Waals
interactions with the internal loop. The global structure of each RNA-small molecule complexes
It is made available under aCC-BY-NC-ND 4.0 International license.
*Author to whom correspondence should be addressed: mdisney@ufl.edu or disney@scripps.edu.
†These authors contributed equally.
SUPPORTING INFORMATION AVAILABLE
(I) TR-FRET analysis of the inhibition of the r(CUG)exp-MBNL1 complex by compounds 1, 2, and 3. (II) H6/H8-H1′ and imino
proton regions of 2D 1H NOESY spectra of unbound r(CUG). (III) Structures of the unbound r(CUG) motif. (IV) 1D 1H and
WaterLOGSY NMR spectra of 1, 2, and 3 alone and in complex with r(CUG). (V) 1D 1H imino region titrations with 1, 2, and 3.
(VI) Imino proton regions of 2D 1H NOESY spectra of the r(CUG)-1, -2, and -3 complexes. (VII) Physicochemical properties of 1, 2
and 3. (VIII) NOE and dihedral restraints used for modeling of the unbound r(CUG) duplex. (IX) NOE restraints used for modeling
of the r(CUG)-1, -2, and -3 complexes. (X) Distance restraint violations greater than 0.1 Å for the r(CUG)-1 complex. (XI) 1H NMR
chemical shifts of the unbound r(CUG) and r(CUG)-1, -2, and -3 complexes. (XII) Synthetic methods for compounds 1 and 2.
HHS Public Access
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retains an A-form conformation, while the internal loops are still dynamic but to a lesser extent
compared to the unbound form. These results aid our understanding of ligand-RNA interactions
and enable structure-based design of small molecules with improved binding affinity for and
biological activity against r(CUG)exp. As the first ever reported structures of RNA r(CUG) repeats
bound to ligands, these structures can enable virtual screening campaigns combined with machine
learning assisted
de novo
design.
INTRODUCTION
The vast majority of the RNA that is transcribed encodes for noncoding (nc)RNAs.1, 2
NcRNAs function in a variety of normal cellular processes, such as splicing, transcription,
translation, degradation, and RNA transport.3 However, these RNAs also cause or contribute
to various diseases, such as cancers, viral diseases, and neurodegenerative diseases,
including those that are caused by RNA repeat expansions.4 RNA repeat expansion diseases
include amyotrophic lateral sclerosis (ALS), Huntington’s Disease (HD), and myotonic
dystrophy types 1 (DM1) and 2 (DM2). DM1 is trinucleotide repeat disorder where an
expanded r(CUG) repeat [r(CUG)exp] is present in the 3′ UTR of the dystrophia myotonia
protein kinase (
DMPK
) gene.4, 5 When the number of repeats exceeds 50,4, 5 the RNA folds
into a long hairpin structure that contains a periodic array of 1×1 UU mismatches.4, 5 These
motifs bind and sequester various RNA-binding proteins (RBPs) such as the alternative pre-
mRNA splicing regulator muscleblind-like splicing regulator 1 (MBNL1).4, 5 The resulting
r(CUG)exp-RBP complexes form foci in the nucleus, leading to poor nucleocytoplasmic
transport of
DMPK
mRNA.6 Loss of functional MBNL1 leads to aberrant pre-mRNA
splicing of its substrates, a hallmark of DM1.4 Huntington’s disease-like 2 is also caused
by r(CUG)exp, which are sequestered into foci containing RBPs.7, 8 The result is loss of
functional JPH3 protein and pre-mRNA splicing defects that cause disease.
Various modalities have been explored as potential treatments for DM1, including antisense
oligonucleotides (ASOs),9 CRISPR/Cas9 gene editing,10 peptides,11 and small molecules.
Several small molecule scaffolds have been reported to bind to r(CUG)exp and inhibit
formation of r(CUG)exp-MBNL1 complexes
in vitro
and in cellular models, including
bis
-
benzimidazole derivatives,6, 12, 13 a triaminotriazine-acridine conjugate,14 pentamidine,15
neomycin B,15 benzo[g]quinoline,16 a perimidine derivative,17 and others.17 To date,
however, there have been no reports of the structures of a small molecule bound to r(CUG)
repeats, whether by X-ray crystallography or NMR spectroscopy, although structures of the
apo form have been reported.18-21 The crystal structure of an RNA is influenced by packing
forces and represent a single state of the RNA.22 In contrast, RNA structures are intrinsically
dynamic and undergo large conformational changes when binding to small molecules.23
While NMR spectroscopy can detect solution state dynamics, NMR spectroscopy of RNA
is often complicated by spectral overlap due to the small chemical shift dispersion of RNA
signals.24 Further, the physicochemical properties of the compound, particularly solubility,
can confound structural studies.
Our laboratory has constructed a database of small molecules that bind to RNA structures,
derived from selection-based experiments or reported in the literature.25, 26 Some of the
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molecules therein elicit effects against their biological targets in patient-derived cells. Three
of these molecules were selected that might be amendable to structure determination using
NMR spectrometry based on their solubilities. Two-dimensional (2D) NMR spectroscopy
and restrained molecular dynamics (MD) were employed to determine the structures of the
complexes formed between all three small molecules and a model of r(CUG) repeats.
METHODS
Calculation of small molecule physicochemical properties.
SwissADME27 was used to calculate physicochemical properties such as molecular weight,
number of hydrogen bond donors and acceptors, number of rotatable bonds and molar
refractivity. Physicochemical properties as well as druglikeness of 1, 2 and 3 showed the
compounds have good druglike properties with no PAINS (pan assay interference structures)
alerts (Table S1).
Small molecule inhibition of r(CUG)n-MBNL1 complex formation in vitro.
A previously reported TR-FRET assay28 was used to study the inhibitory effect of 1-3 on
formation of the r(CUG)12–MBNL1 complex. In summary, 5′-biotinylated r(CUG)12 (80
nM final concentration) was folded in 1× Folding Buffer (20 mM HEPES, pH 7.5, 110 mM
KCl, and 10 mM NaCl) by heating at 60 °C followed by slowly cooling to room temperature
on the benchtop. The buffer was adjusted to 1× Assay Buffer (20 mM HEPES, pH 7.5, 110
mM KCl, 10 mM NaCl, 2 mM MgCl2, 2 mM CaCl2, 5 mM DTT, 0.1% BSA, and 0.5%
Tween-20) and MBNL1-His6 (60 nM final concentration) was then added. The samples were
equilibrated at room temperature for 5 min, and then the compound (1, 2, or 3) was added.
After incubating for 15 min at room temperature, streptavidin-XL665 and anti-His6-Tb
antibody were added to final concentrations of 40 nM and 0.44 ng/μL, respectively, with
a final volume of 10 μL. Samples were incubated for 1 h at room temperature and then
transferred to a 96-well plate. Time-resolved fluorescence was measured on a Molecular
Devices SpectraMax M5 plate reader. Fluorescence was first measured using an excitation
wavelength of 345 nm and an emission wavelength of 545 nm (fluorescence due to Tb).
TR-FRET was then measured by using an excitation wavelength of 345 nm, an emission
wavelength of 665 nm, a 200 μs evolution time, and a 1.5 ms integration time. The ratio of
fluorescence intensity of 545 and 665 nm as compared to the ratios in the absence of ligand
and in the absence of RNA were used to determine IC50s.
Preparation of NMR samples.
The oligoribonucleotide r(GACAGCUGCUGUC) was purchased from Dharmacon and de-
protected per the manufacturer’s recommended protocol. The oligonucleotide was then
de-salted using a PD-10 desalting column containing Sephadex G-25 resin (Cytiva, catalog
#17085101), per the manufacturer’s recommended protocol. After lyophilization to dryness,
the RNA was dissolved in Nanopure water. The concentration of the RNA was determined
by its absorbance at 260 nm, measured at 90 °C using a Beckman Coulter DU800 UV/Vis
spectrophotometer, and the corresponding coextinction coefficient provided by Dharmacon.
This stock was then lyophilized to dryness for NMR experiments to afford the desired
final concentration, 0.3 mM for samples containing unbound RNA or 0.3 mM and 0.4 mM
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for samples containing ligand-bound RNA as indicated. The lyophilized RNA was then
dissolved in NMR Buffer [5 mM KH2PO4/K2HPO4, pH 6.0, and 0.25 mM EDTA] and
folded by heating to 95 °C for 3 min, then slowly cooling the sample to room temperature.
D2O was added to the sample to 5% (v/v) to provide a lock signal. For samples collected
in D2O, the sample was dissolved in NMR Buffer that was exchanged into 100% D2O
and folded by heating to 95 °C for 3 min, and then slowly cooling the sample to room
temperature. To prepare ligand-RNA samples for WaterLOGSY (water-ligand observed via
gradient spectroscopy) experiments, 15 μM of RNA was mixed with 300 μM of compound
to afford a 1:20 RNA:compound ratio.29-31 To prepare ligand-RNA samples for 2D NMR
spectroscopy, 1, 2, and 3 were added to final concentrations of 0.7 mM, 0.6 mM, and 0.6
mM, respectively, where the final concentration of DMSO-d6 was no greater than 5% (v/v).
The concentration of the studied RNA:compound complexes was increased to >300 μM for
2D NMR experiments.
NMR spectroscopy.
NMR spectra of samples in Shigemi tubes (Shigemi, Inc.) were acquired on Bruker Avance
III 600, 700, 850, and 900 MHz spectrometers. 1D and 2D spectra were acquired on the apo
and ligand-bound forms at 5 °C, 6 °C, 15 °C, 25 °C, 30 °C, and 35 °C. NMR experiments
with RNA samples in 95% H2O/5% D2O were carried out at 5 °C, 6 °C, and/or 15 °C. NMR
experiments in with RNA samples in 100% D2O were carried out at 25 °C, 30 °C, and/or
35 °C. Excitation sculpting was applied during acquisition of 1D NMR spectra to suppress
the water signal.32 WaterLOGSY experiments were carried out on samples containing the
r(CUG) duplex construct and each compound.29-31 WaterLOGSY signals were phased to
give negative signals for negative NOEs relative to water. Proton chemical shifts were
referenced to water. 2D 1H-1H NOESY spectra of samples in 95% H2O/5% D2O were
collected with a 125 ms mixing time. 2D 1H-1H NOESY spectra of samples in 100% D2O
were collected with 100, 170, and/or 175 ms mixing times and, separately, at a 400 ms
mixing time. 1D NMR spectra were processed using Bruker TopSpin® and 2D NMR spectra
were processed with TopSpin or NMRPipe33 and assigned with NMRFAM-SPARKY.34
The solubility of small molecules in complex with RNA was measured under NMR
conditions by titrating compound into a solution containing 50 μM of RNA and NMR
Buffer using up to a 6:1 molar ratio of compound:RNA. After addition of each aliquot of
compound, a 1D NMR spectrum was acquired. If the RNA peaks remained visible and
compound peaks appeared in the aromatic region, then a higher concentration sample of the
RNA-compound complex was prepared for 2D NMR spectroscopy.
Methods for obtaining distance and dihedral restraints.
Spectral peaks (NOEs) for 1-, 2-, and 3-bound r(CUG) complexes were assigned using
an NMR solution structure for the apo RNA by comparing spectra of free form and ligand-
bound RNA. Distance restraints for pairs of protons were calculated by integrating NOE
volumes in SPARKY35 or manually assigned a range of distances based on relative NOE
intensities in 100 to 200 ms mixing time NOESY spectra to reduce the influence of spin
diffusion.36 Distance restraints for ligand-RNA NOEs, which were weak in shorter mixing
time spectra, were assigned in the same manner using 400 ms mixing time NOESY spectra.
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NOE volumes were referenced to those calculated from fixed distances: H2′-H1′ (2.75
Å); cytosine or uracil H5-H6 (2.45 Å).37 Hydrogen bonds in canonical AU, GC, and GU
pairs were assigned to distances of 2.1 ± 0.3 Å. NOE intensities between H1′ and H6/H8
indicate that all residues adopted an
anti
conformation. Therefore, the
X
dihedral angle
was constrained between 170° and 340° (
anti
) for all residues except terminal residues and
uridine nucleotides comprising the internal loop in the apo structure.
Modeling methods.
Structures were calculated with a simulated annealing protocol, using the lowest free
energy structure of NMR-restrained apo-r(CUG) structure to which compounds were added
to the binding site in an arbitrary orientation using UCSF Chimera.38 LEaP was then
used to generate topology and coordinate files.39 The starting structure was then energy
minimized using AMBER.39 Restrained molecular dynamics simulations were carried out
with AMBER39 using the parm99X_YIL force field.40 All distance restraints, both inter-
and intra-molecular, for each RNA-small molecule complex can be found in Tables S2 -
S7. Solvation was simulated with the general Born implicit model and 0.1 M NaCl.41 The
system was heated from 0 to 1,000 K in 5 ps, cooled to 100 K in 13 ps, and then to 0 K
in 2 ps. Force constants were 20 kcal mol−1 Å−2 for NOE restraints during the simulated
annealing protocol. The calculation was repeated 100 times. The structures generated from
the calculations were sorted based on NOE distance violations, and the 20 structures with
the fewest violations were selected. The top twenty structures were superimposed and their
RMSD determined using VMD.42
Calculation of global distortions of RNA (curvature).
In order to measure the global deformation of the RNA before and after ligand binding, the
overall bend angle (global curvature) was calculated using
Curves
+.43 The global curvilinear
helical axis was calculated from the average local axes applying a polynomial smoothing.
NMR ensemble with 20 structures was used for this calculation.
RESULTS & DISCUSSION
Compound selection.
To begin to uncover the principles of small molecule recognition of RNA structures, our
laboratory has employed a selection-based method to identify selective RNA motif-small
molecule interactions, which are collected into a database along with interactions reported
in the literature.25, 26 Indeed, these interactions have guided design of molecules that
modulate cellular RNAs in patient-derived cells. Previously, 1 was discovered to bind
the 1×1 nucleotide AA internal loops formed by expanded r(CAG) repeats, or r(CAG)exp
and inhibit complex formation of the RNA with MBNL1,44 the cause of spinocerebellar
ataxia type 3 (SCA3).4 To improve the binding properties of 1 and perhaps direct its
recognition to r(CUG)exp, a derivative of the compound was synthesized, or 2. In 2,
the imidazolines replaced the guanidines of 1 to reduce the polarity of the compound
while substitution of the ester linkage with an amide linkage increases compound stability
and imparts conformational rigidity on the compound.45 Preorganization of the ligand
chemically constrains its structure, reducing the entropic cost of binding to the target
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RNA.46, 47 Compound 3 was a screening hit that bind to the A-bulge present in a mutant
splicing regulatory element (SRE) that controls
MAPT
pre-mRNA alternative splicing, the
presence of which causes frontotemporal dementia with parkinsonism-17 (FTDP-17). The
small molecule was lead optimized by via pharmacophore modeling and chemical similarity
searching.48
1D NMR experiments were then carried out on these compounds to determine if their
solubilities were sufficient under NMR conditions for structure determination. At 50 μM
of RNA and up to 300 μM of compound, resonances for each compound appeared in the
aromatic proton region while imino proton resonances shifted or broadened. Appearance
of the aromatic proton resonances combined with observed changes (broadening and
shifting) of resonances in 1D spectra upon addition of compounds 1 to 3 showed that these
compounds were amenable to structural studies using NMR spectroscopy.
Small molecules 1, 2, and 3 inhibit r(CUG) repeat-MBNL1 complex formation.
The interaction of r(CUG)exp with MBNL1 and other RBPs triggers the formation of nuclear
foci in disease-affected cells.49 Therefore, as a secondary validation of compound binding to
the RNA from 1D NMR spectra, the ability of 1, 2, and 3 to disrupt the r(CUG)exp-MBNL1
complex
in vitro
was measured by using a previously reported time-resolved fluorescence
resonance energy transfer (TR-FRET) assay.28 In this workflow, the RNA–protein complex
was pre-formed followed by addition of the compound of interest at various concentrations
to afford an IC50. The IC50s for 1, 2, and 3 in this assay were 7.9 ± 0.3, 18.9 ± 0.5 and 100.4
± 6.8 μM, respectively (Figure S1). Of note, the affinity of MBNL1 for r(CUG) repeats, for
which the small molecules compete, is 7.2 ± 1 nM.50
NMR analysis of the apo structure of a model of r(CUG) repeats.
Previous NMR and modeling studies with RNA constructs containing one and three tandem
5′CUG/3′GUC motifs show that the 1×1 UU loops are dynamic.21, 51, 52 In this study,
we designed a self-complementary RNA construct (duplex) containing one 5′CUG/3′GUC
motif and five base pairs on either side to stabilize the helical region. The RNA sequence
was optimized to reduce overlap of NMR resonances.
NMR spectra of the free, unbound form of the r(CUG) duplex indicated formation of the
5’CUG/3’GUC internal loop (where the underlined Us indicate loop nucleotides) flanked
by helical regions. In particular, a sequential H6/H8-H1′ walk, in addition to inter-residue
H6/H8-H2′ NOEs, were assigned for all residues in NOESY spectra, indicating that the
duplex adopted an overall A-form structure (Figures S2 - S3 and Tables S2 and S8).
Exchangeable protons were assigned in a spectrum collected in 95% H2O/5% D2O at 5
°C. Cross-peaks between adenine H2 protons and uracil imino protons were assigned for
all AU pairs while cross-peaks between guanine imino and cytosine amino protons were
assigned for all GC pairs except for the terminal GC pair. Intrastrand and interstrand NOEs
from guanine H1 to the 3′ H1′ were observed for the same GC pairs. Specifically, NOEs
were observed from G5H1 to C6H1′ and U10H1′ and G11H1 to U12H1′ and A4H1′ in
both constructs. NOEs from U7H3 to C6H1′, G8H1′, and G8H1, C6H2′ to U7H6, and
U7H1′ and U7H2′ to G8H8 suggest that the loop uridines stack in the helix. This is also
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supported by the observation of the upfield shifted U7H3 proton in the 10-12 ppm range
in the 1D and 2D water experiments, in agreement with chemical shifts of imino protons
in 1×1 UU mismatches.53 These data are consistent with previously reported structures.22,
47, 48 The presence of the U7H3 proton can be used to evaluate the binding of compounds
to the r(CUG) loop. Retention of the U7H3 resonance would be an indicator that the small
molecule binds in a way that does not interfere with the UU mismatch and or stacking.
Loss of the U7H3 resonance would indicate disruption of the UU mismatch with possible
displacement of one or both Us.
Solution structure of the apo RNA.
Structures of apo r(CUG) motif were calculated using 246 distance restraints to generate a
final ensemble of 20 structures (see Methods section). All 20 structures in the final ensemble
adopted an A-form geometry, in agreement with the 2D NMR spectra described above
(Figures 1 and S4 and Table S8). The average RMSD for the ensemble of 20 structures was
1.25 ± 0.33 Å (Table 1), indicating good convergence. The average helical rise and twist
were 2.8 ± 0.3 Å and 30.9 ± 2.0°, respectively. Overall, the helix is slightly under-twisted
compared to A-form RNA, which had an average helical rise and twist of 2.8 Å and 32°,
respectively.21, 54 The average X dihedral angles ranged from −151 to −167° for all residues,
corresponding to
anti
conformations.
Among the 20 ensemble structures, interbase hydrogen bonding of the UU mismatch caused
the helix to be under-twisted at one step (5’CU/3’GU) and over-twisted at the adjacent
step (5’UG/3’UC) within the 5’CUG/3’GUC motif. In particular, the helical twist at the
5′CU/3′GU and 5′UG/3′UC steps were 27.7 ± 11.1° and 33.2 ± 11.7°, respectively. The
UU loop contained a
cis
-Watson Crick/Watson Crick UU pair55 closed by canonical GC
pairs (Figures 1 and S4), stabilized by stacking and hydrogen bonding interactions. The
average number of hydrogen bonds in the UU pair among all 20 ensemble structures was
1.4 ± 0.5. A one hydrogen bonded UU pair was observed in eight of the structures (40%),
and a two hydrogen bonded UU pair was observed in 12 of the structures (60%), indicating
that the loop’s structure is dynamic and interconverting between these hydrogen bonding
structures. Hydrogen bonding results in a small reduction of the average C1′-C1′ distance
for the UU mismatch to 10.0 ± 0.5 Å from an average of ~10.5 Å for A-form RNA, although
the distance is still within range of that observed in A-form RNA.20, 56 The UO4 to UH3
distance was 1.8-1.9 Å in 19 of the 20 structures and >3.4 Å in the remaining structure.
The average buckle, an indication of cusp formation,57 for the UU mismatch was 1.3 ±
6.2°, which is less than values of 5.8 ± 6.4° and −4.2 ± 4.8° calculated for the flanking GC
pairs. The opening, or angle between two bases,57 and stretch, an indication of separation
of two bases,57 of the UU mismatch (−10.3 ± 9.6° and −1.6 ± 0.1°, respectively) were
of the greatest magnitude among all paired nucleotides in the helix. The relatively large
variation for the opening in the UU mismatch is consistent with interconversion of the UU
pair between one and two hydrogen bond states. In summary, the unbound r(CUG) motif
contains a UU pair, stabilized by one or two hydrogen bonds, that induces distortions in the
helix.
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The exchange between one- and two-hydrogen bond states is consistent with that observed
in r(3×CUG).21 The average number of hydrogen bonds in UU pairs calculated by X3DNA
is 1.4 ± 0.5 and 1.5 ± 0.5 for the single r(CUG) and r(3×CUG) constructs, respectively. In
both constructs, hydrogen bonding in UU mismatches was facilitated by shorter C1′-C1′
distances. As in the single r(CUG) construct, the helix is over-twisted and under-twisted at
adjacent steps of the outer 5’CUG/3’GUC motifs in the r(3×CUG). Both base pair steps
of the middle 5′CUG/3′GUC motif of r(3×CUG) are under-twisted; overtwisting instead
occurs at the flanking 5′GC/3′CG motifs. In comparison, Parkesh
et al
. observed exchange
between zero, one, and two hydrogen bond states in their structure of r(1×CUG), with
a shortened C1′-C1′ distance in the two hydrogen bond state. DSSR analysis of their
structures showed that for the one hydrogen bond state, slight undertwisting occurs at the
5′CU/3′GU and 5′UG/3′UC steps and overtwisting occurs at the flanking 5′GC/3′CG
motifs.58 In the two hydrogen bond state, the helical twist (31.5°) at the 5′CU/3′GU
step is close to average but the helix is over-twisted (36.7°) at the 5′UG/3′UC step and
under-twisted at the flanking 5′GC/3′CG steps (28.3° and 25.0°).
Backbone distortion was not observed in a de-twinned crystal structure of r(6×CUG)
that contained a one hydrogen bond stretched UU wobble pair.59 Separately, a construct
containing r(2×CUG) contains a stretched wobble pair, with one hydrogen bond in the
upper 5′CUG/3′GUC motif and two hydrogen bonds in the lower 5′CUG/3′GUC motif. A
shortened C1′-C1′ distance of 8.59 Å was observed in the lower 5′CUG/3′GUC motif to
accommodate the two hydrogen bond state 5’CUG/3’GUC motif.60 In the X-ray structure of
another r(3×CUG) construct, the outer UU mismatches each contained one direct hydrogen
bond, which was the predominant conformation, and the central UU mismatch contained
a water-bridged hydrogen bond.61 The C1′-C1′ distance was 10.8 Å in each of the
outer UU mismatches and 10.3 Å in the central UU mismatch.61 In summary, X-ray and
NMR structures of 5′CUG/3′GUC motifs show that the UU mismatches adopt one or
two hydrogen bond states with helical distortion around the r(CUG) motif. The structural
features of r(CUG) repeats could allow small molecules to recognize and form specific
contacts with r(CUG) motifs.
Analysis of ensembles of NMR structures of fully base paired duplexes using X3DNA62
showed no significant differences in the values of helical parameters compared to those
of apo-r(CUG) or r(3×CUG). The average helical rise and twist values for a dodecamer,
r(CGCAAAUUUGCG)2 (PDB 1AL5) are 2.5 ± 0.6 Å and 33.6 ± 3.1°, respectively.63
For the decamer, r(5′-GAAGAGAAGC/3′CUUCUCUUCG) (PDB 1RRR),64 the average
helical rise and twist are 2.5 ± 0.3 Å and 32.0 ± 2.2°, respectively. All of these values are
within error of those observed for apo-r(CUG) (see above) and r(3×CUG) (2.7 ± 0.5 Å
for helical rise and 30.8 ± 3.7° for helical twist).21 For the decamer, r(5′-GCAGAGAGCG/
3′-CGUCUCUCGC) (PDB 2JXQ),65 the average helical rise and twist are 2.4 ± 0.3 Å
and 33.7 ± 2.4°, respectively. The average C1′-C1′ distance for each base pair was in
the 10.7-10.8 Å range in PDB 1AL5,63 and the 10.7-10.9 Å range for PDB 1RRR,64
and 10.4-10.6 Å for PDB 2JXQ, which as expected for A-form RNA.65 As noted earlier,
r(CUG)-containing RNAs typically have shorter C1′-C1′ distances, particularly for two
hydrogen bond UU pairs. Average major groove width values ranged from 18.4-19.7 Å for
PDB 1AL5, 19.3-21.8 Å for PDB 1RRR, and 16.4-16.9 Å for PDB 2JXQ. In comparison,
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these values ranged from 18.9-19.6 Å for apo-r(CUG) and from 17.8-20.6 Å for r(3×CUG).
Thus, the helical structures of apo-r(CUG) and r(3×CUG) appear to be consistent with those
of fully base paired duplexes. That said, larger variations in the C1′-C1′ distances and
number of hydrogen bonds are observed for mismatched UU pairs in the apo-r(CUG) and
r(3×CUG) RNAs than for canonical base pairs in the same constructs and the fully base
paired constructs. Thus, small molecules could selectively recognize the localized dynamics
in the UU pairs, rather than the helical structure it shares with fully paired RNAs, and form
favorable interactions that compete with MBNL1 binding.
NMR analysis of 1D and WaterLOGSY spectra.
As a first step, the molecules were studied in a WaterLOGSY experiment. WaterLOGSY
spectra of 1, 2, and 3 in the absence of RNA gave positive NOEs, indicating no aggregation
of the compounds. Addition of r(CUG) duplex to each compound gave negative NOEs
(positive WaterLOGSY signals) for the compounds (Figures S5 - S7). WaterLOGSY spectra
of 1, 2, and 3 in the presence of RNA also gave RNA signals with negative intensity.
Thus, these negative NOEs resulted from small molecule binding to the RNA, rather than
aggregation of the compounds. Thus, these spectra indicate that the RNA-small molecule
complexes were soluble.
Complementarily, imino proton spectra of the duplex model were collected as a function of
small molecule concentration. For 1, addition of compound to 1:1 1:RNA ratio caused minor
shifting of the U10H3 and G5H1 resonances, suggesting that the compound is binding to the
r(CUG) motif (Figure S8). At this ratio, U7 broadened significantly while G8H1 was mostly
unchanged. At 4:1 1:RNA, U7H3 was broadened while G8H1 continued to shift upfield,
indicating improved stacking interactions in the r(CUG) motif and slight displacement of the
uracils from the helix. At this ratio, G1H1 also began to shift upfield, indicating stacking
of the compound at the end of the helix. Addition of the small molecule to 6:1 1:RNA
molar ratio caused U10H3 to shift further downfield and U12H3 and G5H1 to shift upfield,
respectively, relative to the unbound state. Within the CUG motif, G8H1 continued to shift
upfield from its position at the 4:1 1:RNA ratio and broadened. Although these shifts may
be minor, the shifting and broadening of peaks, particularly around the r(CUG) motif,
collectively indicate a structural shift upon compound binding. These results also show that
the compound selectively binds to r(CUG) over base paired regions.
Addition of 2 to 2:1 2:RNA resulted in broadening of G5H1, U7H3 and G8H1, but the
resonances did not otherwise shift. At 4:1 2:RNA, U7H3 and G8H1 broadened further and
G8H1 shifts downfield, towards G5H1, indicating binding of the compound to the r(CUG)
motif. Meanwhile, U10H3 shifted downfield relative to the unbound duplex. At 6:1 2:RNA
molar ratio, addition of 2 resulted in a further downfield shift of U10H3 relative to unbound
but G5H1, U7H3, and G8H1 do not further broaden. This indicates saturation of compound
at the r(CUG) motif at the 4:1 2:RNA molar ratio. G1H1 and G11H1 remained overlapped
upon addition of compound to 6:1 2:RNA ratio (Figure S9). The shifting and broadening of
peaks suggest that the compound binds to the r(CUG) motif.
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Addition of 3 at a 1:1 ratio of compound:RNA shifted G5H1 and G8H1 upfield and U7H3
and U10H3 downfield. At 4:1 3:RNA ratio, G5H1 shifted further downfield, G8H1 shifted
further upfield, and U7H3 broadened. This suggests that 3 disrupts the UU base pair at this
ratio. At 6:1 compound:RNA ratio, G8H1 continued to shift upfield while U7H3 remained
broadened. Otherwise, resonances in the r(CUG) motif do not differ much at the 4:1 and
6:1 1:RNA ratios, indicating saturation of binding at that site. Away from the r(CUG) motif,
shifts in G11H1 and U12H3 were less pronounced throughout the titration. However, G1H1
is distinguishable from G11H1 4:1 3:RNA ratio, which may result from stacking of the
compound at the end of the helix (Figure S10).
Collectively, both the WaterLOGSY and imino proton spectra suggest that the small
molecule binds to the RNA with sufficient affinity and selectivity to determine the structures
of the complexes.
NMR analysis of r(CUG) bound to 1 and 2.
As discussed above, an uninterrupted NOESY walk with sharp cross-peaks was observed for
the apo form of the r(CUG) repeat duplex model. Interestingly, an uninterrupted NOESY
walk was also observed in the spectra collected for the duplex in the presence of related
small molecules 1 and 2 (Figures 2 - 3 and S11 - S12). Further, these cross-peaks remained
sharp (similar to those observed in the apo spectra), suggesting fast exchange among
multiple states of the complexes. A total of 24 NOEs between the RNA and 1 were observed
for the r(CUG)-1 complex (Figure 2 and Table S9). Among them, NOEs were observed from
H1 and H3 aromatic protons of 1 to sugar protons of C6, and G8 and to aromatic protons
of C6, U7, and G8. On the opposite side of the 1, NOEs were observed from H5 and H7
aromatic protons of 1 to aromatic and sugar protons of U20 and G21. However, no NOEs
between 1 and the loop uracils were observed, Thus, 1 appears to bind to the major groove
of the RNA, between the closing GC pairs of r(CUG) motif and disrupts base pairing and/or
stacking interactions of the UU mismatch.
In the r(CUG)-2 complex, 23 NOEs between r(CUG) and 2 were observed (Figure 3
and Table S10), six of which are also observed between r(CUG) and structurally similar
compound 1: H1 of 2 to U7H5 and U7H6; H5 of 2 to C19H2′, U20H5, U20H6; H7 of 2
to U20H5. In addition to these and other NOEs between 2 and sugar and aromatic protons
of r(CUG), 2 has NOEs from the aromatic H4 and H6 protons of 2 to U7H3 and U20H3,
respectively, from one of the imidazoline groups of 2 to U20H6 and G21H5”, and from
the H26 amide proton of 2 to U7H3. These NOEs between the compound and the imino
protons of the loop uracils laid within the 10-12 ppm imino proton range, where U7/U20H3
of the apo structure resonates, suggesting that the loop uracils remain base paired. That
said, broadening of the U7 and G8 imino protons upon addition of 2 as observed in the 1D
titration indicates that the complex is dynamic. In summary, the NMR data suggests that 2
adopts a similar binding mode to 1, where the compound laid in the major groove of the
RNA and between the closing GC pairs of r(CUG), but with a different conformation of the
UU mismatch. As with 1, each of the aromatic rings appeared to be in close proximity to
one, but not both strands of the RNA.
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Structures of r(CUG) repeat-1 and r(CUG) repeat-2 complexes.
As expected from the uninterrupted NOESY walk observed for both small molecules
(Figures 2 - 3), the r(CUG) RNA retained A-form geometry when bound to 1 or 2. A total
of 24 and 23 intermolecular NOEs were used as distance restraints to model the r(CUG)-1
and -2 complexes, respectively (Tables 1, S4 - S6). The average RMSD of all heavy atoms
for the ensemble of structures from restrained MD was 1.81 ± 0.43 Å for r(CUG)-1 and 1.87
± 0.48 for r(CUG)-2, both slightly higher than that of the apo structure (1.25 ± 0.33 Å; Table
1).
Although small molecules 1 and 2 both laid in the major groove of the 5’CUG/3’GUC
internal loop motif, different intermolecular interactions afford stability to the complexes
(Figures 1 and 4). Small molecule 1 formed stacking interactions with the closing GC
pair(s), but not the uracil loops. Both uracil loops were stacked in the helix in 19 of
the 20 structures; in the remaining structure one uracil was flipped into the major groove
and formed one hydrogen bond with a guanidine of 1. However, the uracils were shifted
away from each other in base pair plane, disrupting their interbase hydrogen bonding
interactions. Additionally, they did not form contacts with 1, consistent with the absence
of NOEs between 1 and the uracils. The positively charged guanidines formed electrostatic
interactions (5 Å distance or less)66, 67 with the phosphate backbone at G8 and G21. In all
structures that comprise the ensemble formed by the r(CUG) repeat-1 complex, the carbonyl
oxygen of 1 was oriented towards the minor groove. The major and minor grooves within
the 5’CUG/3’GUC motif were widened in the r(CUG)-1 structures, and the loop’s closing
GC pairs had the largest changes in buckle of the C6G21 (14.2 ± 6.7° vs. 5.8 ± 6.4° in the
apo structure) and G8C19 pairs (−15.8 ± 7.1° vs. −4.2 ± 4.8° in the apo structure).
Not surprisingly, addition of 1 to r(CUG) disrupted hydrogen bonding within the UU
mismatch and resulted in increases in helical twist at 5′CU/3′GU and 5′UG/3′UC steps
within the 5’CUG/3’GUC motif. At one of the 5′GC/3′CG steps, the helical twist increases
from 31.7 ± 1.5° to 35.3 ± 3.8° while at the other 5′GC/3′CG step, the helical twist remains
relatively unchanged (33.2 ± 11.7°) in the apo structure and 33.5 ± 3.1° in the 1-bound
structure). Changes in values of 5′GC/3′CG parameters also suggest that helical regions
upstream of the UU mismatch undergo rearrangement to accommodate binding of the
compound. The bend angle of the helix decreased from 11.88 ± 5.01° for the apo structure
to 9.53 ± 4.26° for the 1-bound structure. The major groove width at the 5′CU/3′GU
and 5′UG/3′UC steps and flanking 5′GC/3′CG steps increase upon binding of 1. These
results suggest that the compound affects the helix away from the r(CUG) motif, consistent
with shifting of imino proton resonances in base pairs away from the r(CUG) motif. Large
variation in these values suggest that the r(CUG) motif in the apo and bound structures are
dynamic and that binding of 1 disrupts localized interactions that stabilize the r(CUG) motif.
The overall average helical rise and twist of all nucleotides bound RNA were similar to that
of the apo structure, 2.2 ± 1.0 Å and 31.8 ± 3.0° (2.8 ± 0.3 Å and 30.9 ± 2.0°, respectively,
for the apo structure). However, the increased errors in these values as compared to those
of apo-r(CUG) suggest that the r(CUG)-1 complex is dynamic. An increase in the C1′-C1′
distance at the UU mismatch in the bound structure was also observed, as compared to the
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apo structure, 12.8 ± 0.7 Å and 10.0 ± 0.5 Å, respectively. Large propeller twist values
were also observed for the UU pair in the r(CUG)-1 complex and were significantly larger
than observed in the unbound r(CUG) repeat structure. However, the values for shear (a
measure of sliding of one base in relation to the other)57 were not significantly different
between the two structures; rather, base non-planarity is represented in the stagger values of
the r(CUG) repeat-1 structures. Altogether, the r(CUG)-1 complex is stabilized by stacking
and hydrogen bonding interactions and remodels the RNA structure around the UU loop, by
decreasing the bend angle of the helix, increasing the major groove width within the r(CUG)
motif, disrupting hydrogen bonding in the UU mismatch, and lengthening the C1′-C1′
distance of the UU mismatch.
As mentioned above, the binding of 2 to the r(CUG) repeat did not disrupt the overall
A-form geometry of the RNA, as evidenced by the average helical rise (2.8 ± 0.5 Å) and
helical twist (30.6 ± 19.9°) for the ensemble of 20 r(CUG) repeat-2 structures generated
by restrained MD (Figure 1). Although these values are similar for the apo structure, the
large error in helical twist suggests that the r(CUG)-2 complex is dynamic. Although 2 also
formed stacking interactions the 5’CUG/3’GUC internal loop motif as 1 did, 2 stacked on
the mismatched uracils, rather than the GC closing base pair (Figure 4). In further contrast
to 1, in most of the r(CUG)-2 structures (14/20), either one or both UU mismatch bases
were not stacked in the helix and displaced towards the minor groove. However, the uracils
remained base paired, as evident from an average of 1.6 ± 0.5 hydrogen bonds vs. 0.2 ±
0.4 for the U7U20 pair in the 1-bound structure. The mismatched uracils were also in close
proximity to 2, consistent with NOEs between U7/U20 imino proton and 2. The interactions
between 2 and the UU loop were also stabilized by a hydrogen bond (3 Å distance or
less) between an uracil of 5’CUG/3’GUC internal loop and the amide H26 of the small
molecule (12/20 structures) and a hydrogen bond between the phosphate backbone at U20
and H27 of 2 (16/20 structures). The imidazolines form electrostatic interactions with the
phosphate backbone at C6 in 10 of the 20 structures, at U7 in all of the structures, at
U20 in 19 of the 20 structures, and at G21 in six of the 20 structures. Aromatic protons
of the benzylimidazoles formed van der Waals and/or nonconventional hydrogen bonding
interactions with G8, C19, and/or G21G21.
A closer inspection of the structure of the internal loop bound to 2 indicates that the
5′CU/3′GU step is under-twisted and the 5′UG/3′UC step is over-twisted, concomitant
with a much shorter C1′-C1′ distance, 9.6 ± 1.0 Å. Farther away from the r(CUG) motif,
the 5′GC/3′CG steps appear to be over-twisted. This under- and over-twisting and the
shortening of C1′-C1′ distance is more severe in the r(CUG)-2 complex than that observed
in both apo structure or in the structure of 1 complex. Akin to that observed in the structure
of the r(CUG) repeat-1 complex, a large propeller twist value, 25.8 ± 29.0°, was observed
for the UU pair in the r(CUG) repeat-2 structure vs. −9.8 ± 3.6° in the unbound structure
while the shear values were not significantly different. Rather, stagger values for r(CUG)
repeat-2 structures (−1.1 ± 1.8° for C6G21) indicated non-planarity of the RNA bases.
Collectively, these changes upon 2 binding of the internal loop slightly widened. The large
error values in the parameters for the 2-bound structure indicate that the complex is dynamic
as the helix accommodates binding of the compound. The dynamics observed in the r(CUG)
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motif are also evident from breathing of the closing GC pairs, which have average numbers
of hydrogen bonds of 2.2 ± 1.3 for the C6-G21 and 2.8 ± 0.5 for the G8-C19 pair. The
changes in the structure of r(CUG) are also consistent with shifting of imino resonances
upon titration of 2 into r(CUG). In summary, the binding of 2 to r(CUG) repeat distorts
the RNA helix by increasing the bend angle by 2.68° (average of 14.81 ± 2.68°) and
compensates for those distortions by forming intermolecular stacking and hydrogen bonds.
Comparison of the r(CUG)-1 and r(CUG)-2 structures.
The Carloni group predicted the structure of 1 bound to r(CAG) using docking and
molecular dynamics simulations.68 In their structures, 1 binds to the major groove of r(CAG)
and interacts with both strands of the RNA. The hydrogen bonding interactions of one of
the AA pairs (A7-A14) appears to be altered from the free state when bound to 1. In the
r(CUG)-1 and -2 NMR structures, the compounds also bind in the major groove, but 1
disrupted the hydrogen bonding interactions in the UU mismatch whereas 2 did not. In the
Carloni group’s structure of 1 bound to a r(CAG) repeat, hydrogen bonds were observed
between the guanidines of 1 and the phosphate backbone of the RNA. In the r(CUG)-1
and r(CUG)-2 structures, the guanidines of 1 and imidazolines of 2 also formed hydrogen
bonds and electrostatic interactions with the negatively charged phosphates. The modeled
r(CAG)-1 structures have stacking interactions between the rings of 1 and the mismatched
bases of the 1×1 AA pairs. In comparison, stacking interactions between the compound
and mismatched uracils were observed in the r(CUG)-2 but not -1 structures. To facilitate
these stacking interactions, the rings in 2 adopted a quasi-perpendicular orientation in some
and a planar orientation in the other states. The quasi-perpendicular orientation of the rings
in 1 was present in the Carloni group’s r(CAG)-1 structures, but not the NMR structures
of r(CUG)-1, where the rings adopted a planar orientation in all states. The mismatched
uracils in the r(CUG)-1 structures formed stacking interactions with the helix, but lost their
interbase hydrogen bonding, whereas those in the r(CUG)-2 structures retained interbase
hydrogen bonding but lost their stacking interactions with the helix. The mismatched A4-
A17 and A7-A14 bases in the Carloni group’s structures and the U7-U20 bases in the
r(CUG)-1 structures had negative opening values, but the buckle values of both closing GC
pairs were negative in the r(CAG)-1 structure and alternated between negative and positive
in the r(CUG)-1 structures. In the r(CUG)-2 structures, the opening was positive and the
average buckle values did not change significantly from apo-r(CUG). The major groove
widened upon binding of compound in the Carloni group’s work and in the r(CUG)-1 and
-2 structures. Altogether, binding of 1 and 2 to r(CUG) and r(CAG) induces changes in the
local structure of the RNAs but differentially affects interactions between the mismatched
bases. Furthermore, selectivity of the compounds for r(CUG) motifs can be increased by
designing the compounds to interact with the UU mismatches and closing GC pairs.
NMR analysis of r(CUG) bound to 3.
As observed for 1 and 2, binding of 3 did not disrupt a NOESY walk through the r(CUG)
repeat duplex model (Figure 5), indicating that an overall A-form geometry was adopted. In
total, 32 NOEs were observed from 3 to the RNA (Figure 5 and Table S11), more than that
observed for 1 (24) or 2 (23). NOEs were also observed between H5, H6, and H8 of 3 and
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the RNA, suggesting that the methoxybenzene group rotates about the C10-N3 bond upon
binding to the RNA (Figures 5 and S13). Of note, the NOEs between 3’s H5 and the r(CUG)
motif were not used as restraints for modeling because they were close to the level of noise
in the 100 ms mixing time NOESY spectrum. The average RMSD for the 20 structures with
the fewest distance restraint violations was 1.02 ± 0.20 Å (Table 1).
Structure of the r(CUG) repeat-3 complex.
As aforementioned, the overall r(CUG) repeat-3 complex formed an overall A-form
geometry as confirmed by its average helical rise and twist for the ensemble structures,
2.5 ± 0.6 Å and 32.8 ± 11.1°, respectively (Figure 1). The large error in helical twist
suggests that the complex is dynamic. Perhaps unexpectedly, the binding mode for 3 and
its subsequent effect on the RNA’s structure is markedly different than that of 1 and 2,
where stacking interactions, hydrogen bonds, and van der Waals interactions stabilize the
r(CUG)-3 complex (Figure 4). The quinoline and methoxybenzene groups of 3 stacked
between the closing GC pairs of the 5’CUG/3’GUC motif and displaced one of the uridines
of the UU mismatch into the major groove and the other into the minor groove. (Small
molecule 1 also stacked on the GC pairs.) The C7 methyl group was oriented towards the
minor groove while the C15 methyl group pointed towards the phosphate backbone of the
RNA. These groups formed non-conventional hydrogen bonds / van der Waals interactions
with the RNA. These types of interactions between the small molecule’s H3 and H7 and the
loop nucleotides were also common in the ensemble of 20 r(CUG)-3 structures. H13 and
H14 of the compound formed hydrogen bonds with cytidines forming the closing GC pairs
on the opposite strand of the RNA and the amidine lied inside the major groove and forms
electrostatic interactions with the phosphate backbone at C6. Taken together, the r(CUG)-3
complex is stabilized by a combination of stacking, van der Waals, and hydrogen bonding
interactions.
Although the overall average twisting of the r(CUG) repeat-3 complex indicated A-form
geometry, the helix appears to be under-twisted at the 5′CU/3′GU step and over-twisted at
the 5′UG/3′UC step. At the 5′GC/3′CG steps adjacent to the 5’CUG/3’GUC internal loop,
the helix is over-twisted (39.8 ± 2.1° and 35.2 ± 0.9°) while the bend angle of the helix
increased to 13.92 ± 5.06°. To accommodate 3, the major groove was slightly narrower at
the 5′CU/3′GU and 5′UG/3′UC steps (17.8 ± 0.9 Å and 17.7 ± 0.9 Å, respectively) and at
one of the 5′GC/3′CG steps (17.4 ± 0.7 Å) than that observed for the same steps) in the apo
structure (19.6 ± 1.2 Å for 5′CU/3′GU and 5′UG/3′UC steps and 19.6 ± 1.0 Å for the same
5′GC/3′CG step). The changes in the 5′GC/3′CG steps were consistent with shifts in G5H1
and G8H1 resonances in the titration experiment. An increase in C1′-C1′ distance from 10.0
± 0.5 Å in the unbound structure to 14.6 ± 0.2 Å disrupted the UU base pair and allowed
the compound to stack between the closing GC pairs. Nonplanarity of the UU mismatch was
also evident from high values for parameters such as buckle (22.5 ± 60.3°), shear (14.5 ±
0.8°) and stagger (4.8 ± 1.5°) for the UU pair, and changes in rise at the 5′CU/3′GU (5.9
± 0.9 Å vs. 3.7 ± 0.2 Å in apo structure) and 5′UG/3′UC steps (−0.8 ± 1.5 Å vs. 3.7 ±
0.2 Å in the apo structure), roll at the 5′CU/3′GU (−41.5 ± 21.3° vs. 9.2 ± 3.8° in the apo
structure) and 5′UG/3′UC steps (13.4 ± 7.5° vs. 7.3 ± 2.5° in the apo structure), and tilt at
the 5′CU/3′GU (−73.4 ± 39.5° vs. 0.6 ± 3.4° in the apo structure) and 5′UG/3′UC steps
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(83.3 ± 43.7° vs. −2.7 ± 3.4° in the apo structure). The large errors in values for the base pair
parameters are consistent with dynamics observed in the ensemble of r(CUG)-3 structures.
In summary, binding of 3 to r(CUG) brings about structural changes and dynamics in the
helix.
Comparison of the r(CUG)-3 structure with tau RNA-3 structure.
In the r(CUG)-3 structures and tau RNA-3 structures, 3 lied in the major groove of the
RNA and stacks with the RNA via its quinoline group. The C7 methyl group in these
structures was positioned between the closing GC pairs of the A-bulge or UU mismatch and
forms van der Waals and/or nonconventional hydrogen bonds with the closing GC pair(s).
Similarly, the C15 group interacted with the RNA backbone in the complexes. Compared
to the tau RNA-3 structures, the quinoline core in the r(CUG)-3 structures is turned slightly
towards one of the RNA strands. This orients the amidine away from the RNA backbone
of one of the RNA strands and positions the N3 amino group closer to the opposite strand
and facilitates a hydrogen bond with the C19 base. Taken together, the r(CUG)-3 complex
resembles the tau RNA-3 complex and is stabilized by a similar set of interactions.
Comparison of the r(CUG) repeat structures bound to 3 vs. 1 and 2.
The r(CUG)-3 complex, like r(CUG)-1 and r(CUG)-2 complexes, contained the compound
in the major groove between the closing GC pairs, with the helices retaining A-form
geometry. Whereas 1 and 2 formed stacking interactions with either of the closing GC
pairs or the UU mismatch, 3 stacked with both of the GC pairs. To allow for this, the
helix was bent to a greater degree in the r(CUG)-3 structures than the apo or r(CUG)-1
structures, although to a similar degree as the r(CUG)-2 structures as a result of stacking
of the extrahelical UU pair in most of the r(CUG)-2 conformations. That said, one of the
uracils in the r(CUG)-3 structures remained in the helix and stacked on the compound
while the other was flipped into the major groove. As a result, the C1′-C1′ distance
of the mismatched uracils in the r(CUG)-3 was greater than in r(CUG)-1 and r(CUG)-2
structures. Compound 3 also contained methyl groups positioned near the RNA backbone
for weak nonconventional hydrogen bonds or van der Waals interactions with the RNA. In
comparison, the imidazolines of 2 laid farther in the major groove and did not interact with
the RNA. Instead, 2 formed a more extensive hydrogen bonding network with the RNA than
1 or 3. Altogether, 3 appears to form more stabilizing interactions with r(CUG) than 1 or
2. In summary, compounds 1, 2, and 3 form diverse sets of interactions that aid molecular
recognition of r(CUG) repeats.).
Implications & Conclusions.
Structure-based drug design is powerful tool for drug discovery as evidenced by various
efforts to identify and optimize small molecules that bind disease-causing proteins.69
Unfortunately, such methods are only in their infancy for RNA targets. Herein, we
used the physicochemical properties of known small molecules that bind RNA, coupled
with solubility measurements to identify three small molecules that bind r(CUG) repeats,
amenable to structure determination by NMR spectrometry. NMR spectrometry, has several
advantages over other structure determination techniques such as X-ray crystallography
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and cryo-EM,70 particularly its ability to capture conformational dynamics of the target
and target-ligand complexes. Indeed, conformational dynamics have been incorporated in a
virtual screen to find small molecules that bind HIV TAR RNA.23
This study provides the very first examples of small molecules bound to the UU internal
loops found in r(CUG)exp. Compound 1 was previously identified to bind to 1×1 AA loops
and inhibit MBNL1 sequestration in HD.44 For this reason, it was used to initiate structural
studies of r(CUG)exp-binding compounds that inhibit MBNL1 sequestration in HDL2. Using
the NMR structure of r(CUG)-1, 2 was designed with improved chemical properties and
a structure of the compound in complex with r(CUG) was solved. Separately, an NMR
structure was determined for r(CUG) in complex with 3, which was previously identified to
bind to an A-bulge and inhibit U1 snRNP binding in FTDP-17.48 These compounds each
bound to the r(CUG) internal loop via unique sets of interactions. Thus, a structure-based
approach to identify RNA-binding compounds may be initiated by searching for compounds
that bind to RNA motifs, testing their solubility under NMR conditions, and then analyzing
chemical shift perturbations of the RNA upon binding of the compounds.
The selectivity of 1 – 3 for r(CUG)exp could be improved by optimizing their interactions
with the 5’CUG/3’GUC internal loop, such as by eliminating steric hindrance or adding new
hydrogen bonding and van der Waals interactions. This approach has been used to optimize
small molecule drugs that target proteins.71 Compounds that selectively bind to RNA can
also be identified by docking libraries of compounds to ensembles of RNA conformations
generated by NMR studies and computational modeling.23, 72 Using ligand-bound receptors
usually affords superior results than apo receptors in molecular docking because the binding
sites are better defined in the bound structures than unbound ones.73 Thus, the results of
this work provide invaluable information for future virtual screening campaigns against
compounds that target r(CUG)exp. These data can also be used for machine learning based
methods to improve the binding affinity as well as physicochemical properties through
approaches like generative artificial intelligence (AI).74 These structures also provide the
opportunity to adopt structure-based machine learning assisted drug design approaches as
already exemplified in the protein-small molecule field.75, 76
Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
ACKNOWLEDGMENTS
This work was funded by the National Institutes of Health (R35 NS116846 to M.D.D), the Department of Defense
(HT94252310336 to M.D.D), the Muscular Dystrophy Association (MDA 1069959 to M.D.D and Development
Grant 22 grant 963835 to A.T.), the Huntington’s Disease Society of America (to J.L.C.). This study made use of
NMR spectrometers at The Scripps Research Institute, Campus Chemical Instrument Center NMR facility at Ohio
State University, Minnesota NMR Center, and the National Magnetic Resonance Facility at Madison (NMRFAM).
NMR instrumentation at The Scripps Research Institute was purchased with funds from the NIH S10-OD021550.
NMR instrumentation, helium recovery equipment, and computers at NMRFAM were purchased with funds from
the University of Wisconsin-Madison, the NIH P41GM136463, R24 GM141526, P41 GM103399, S10 RR023438,
S10 RR025062, and S10 RR029220.
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ABBREVIATIONS
1D one-dimensional
2D two-dimensional
ALS amyotrophic lateral sclerosis
ASO antisense oligonucleotide
CRISPR clustered regularly interspaced palindromic repeats
CUGBP1 CUG-binding protein 1
DM1 myotonic dystrophy type 1
DMPK dystrophia myotonica protein kinase
FTDP-17 frontotemporal dementia with parkinsonism-17
HD Huntington’s disease
HDL2 Huntington’s disease-like 2
IR insulin receptor
MBNL1 muscleblind-like 1 protein
MD molecular dynamics
nc noncoding
NMR nuclear magnetic resonance
NOE nuclear Overhauser effect
NOESY nuclear Overhauser effect spectroscopy
RBP RNA-binding protein
RMSD root-mean-square deviation
SCA spinocerebellar ataxia
SRSF6 serine and arginine rich splicing factor 6
UTR untranslated region
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Figure 1: NMR solution structures of unbound r(CUG) duplex and r(CUG) duplex bound to
compounds 1-3.
(A) Structure of unbound r(CUG). (B) Structure of the r(CUG)-1 complex. (C) Structure of
the r(CUG)-2 complex. (D) Structure of the r(CAG)-3 complex. For each model, an overlay
of the 10 structures with the fewest distance restraint violations in stick representation (left)
and a surface representation of the structure with the fewest distance restraint violations
(right) are shown.
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Figure 2: H6/H8-H1′ region of a 2D 1H-1H NOESY spectrum of the r(CUG)-1 complex.
Blue lines represent a sequential H6/H8-H1′ walk, and blue labels represent intraresidue
H6/H8 to H1′ NOEs. Dashed red lines represent adenine H2 resonances, and red labels
represent interresidue and intraresidue NOEs between adenine H2 and H1′ of nearby
residues. Dashed light blue lines represent aromatic proton resonances of 1, and light blue
labels correspond to NOEs between H1′ of r(CUG) residues, unless otherwise noted, and 1.
RNA residues are only numbered from 1 to 13 in the NMR spectrum, and residues 14 to 26
correspond to residues 1 to 13 due to the symmetry of the RNA construct. In the chemical
structure of 1, black numbers represent hydrogens, gray numbers represent carbons, blue
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numbers represent nitrogens, and red numbers represent oxygen. NOEs between the RNA
and 1 and their corresponding distance restraints used for modeling are colored according to
the atoms of 1. The spectrum was acquired at 25 °C with 400 ms mixing time and 0.3 mM of
RNA and 0.6 mM of 1.
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Figure 3: H6/H8-H1′ region of a 2D 1H-1H NOESY spectrum of the r(CUG)-2 complex.
Blue lines represent a sequential H6/H8-H1′ walk, and blue labels represent intraresidue
H6/H8 to H1′ NOEs. Dashed red lines represent adenine H2 resonances, and red labels
represent interresidue and intraresidue NOEs between adenine H2 and H1′ of nearby
residues. Dashed light blue lines represent aromatic proton resonances of 2, and light blue
labels correspond to NOEs between H1′of r(CUG) residues, unless otherwise noted, and 2.
RNA residues are only numbered from 1 to 13 in the NMR spectrum, and residues 14 to 26
correspond to residues 1 to 13 due to the symmetry of the RNA construct. In the chemical
structure of 2, black numbers represent hydrogens, gray numbers represent carbons, blue
numbers represent nitrogens, and red numbers represent oxygen. NOEs between the RNA
and 2 and their corresponding distance restraints used for modeling are colored according to
the atoms of 2. The spectrum was acquired at 35 °C with 400 ms mixing time and 0.3 mM of
RNA and 0.6 mM of 2.
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Figure 4: 3D and schematic diagrams of ligand-r(CUG) interactions.
(A) Interactions in the r(CUG)-1 complex. (B) Interactions in the r(CUG)-2 complex. (C)
Interactions in the r(CUG)-3 complex. In the 3D and schematic diagrams, light green dashed
lines represent conventional hydrogen bonds, purple dashed lines represent electrostatic
interactions, magenta dashed lines represent nonconventional hydrogen bonds, and light
blue dashed lines represent stacking interactions. In the schematic diagrams, dark blue lines
represent van der Waals interactions (not shown in the 3D diagrams).
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Figure 5: H6/H8-H1′ region of a 2D 1H-1H NOESY spectrum of the r(CUG)-3 complex.
Blue lines represent a sequential H6/H8-H1′ walk, and blue labels represent intraresidue
H6/H8 to H1′ NOEs. Dashed red lines represent adenine H2 resonances, and red labels
represent interresidue and intraresidue NOEs between adenine H2 and H1′ of nearby
residues. Dashed light blue lines represent aromatic proton resonances of 3. Light blue
labels around the edges of the spectrum correspond to r(CUG) and aromatic hydrogens of
3 in RNA-3 NOEs. RNA residues are only numbered from 1 to 13 in the NMR spectrum,
and residues 14 to 26 correspond to residues 1 to 13 due to the symmetry of the RNA
construct. In the chemical structure of 3, black numbers represent hydrogens, gray numbers
represent carbons, blue numbers represent nitrogens, and red numbers represent oxygen.
NOEs between the RNA and 3 and their corresponding distance restraints used for modeling
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are colored according to the atoms of 3. The spectrum was acquired at 35 °C with 400 ms
mixing time and 0.4 mM of RNA and 0.6 mM of 3.
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Table 1:
Structural refinement statistics for the average of 20 structures for unbound and ligand bound RNAs.
Unbound
r(CUG) r(CUG)-1 r(CUG)-2 r(CUG)-3
No. of restraints
All NOE restraints and hydrogen bonds 246 336 292 358
All NOE restraints 214 284 266 326
Intraresidue/compound intramolecular 118 134 122 161
Sequential residues 88 126 104 133
Long range, excluding RNA-compound 8 0 17 5
RNA-compound N/A 24 23 27
Hydrogen bond 32 52 26 32
Dihedral/chiral restraints 130 0 0 0
RMSD of experimental restraints
Distance (Å) 7.9 × 10−4 1.0 × 10−3 9.3 × 10−4 2.8 × 10−3
Dihedral (deg) 0.3 0.0 0.0 0.0
RMSD of structures for heavy atoms (Å)
All residues, excluding compound 1.25 ± 0.33 1.81 ± 0.43 1.87 ± 0.48 1.01 ± 0.19
All residues and compound N/A 1.86 ± 0.42 1.89 ± 0.48 1.02 ± 0.20
Outer helices (residues 1-5 and 9-13) 1.22 ± 0.36 1.88 ± 0.49 1.75 ± 0.53 0.93 ± 0.22
Compound N/A 1.02 ± 0.25 0.94 ± 0.30 0.46 ± 0.29
Internal loop and closing pairs (residues 6-8), excluding compound 0.78 ± 0.30 0.94 ± 0.28 1.65 ± 0.62 0.79 ± 0.18
Internal loop and closing pairs (residues 6-8) and compound N/A 1.18 ± 0.31 1.65 ± 0.59 0.80 ± 0.15
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