SiRA: A Silicon Rhodamine-Binding Aptamer for Live-Cell Super-Resolution RNA Imaging

Article abstract of DOI:10.1021/jacs.9b02697

Although genetically encoded light-up RNA aptamers have become promising tools for visualizing and tracking RNAs in living cells, aptamer/ligand pairs that emit in the far-red and near-infrared (NIR) regions are still rare. In this work, we developed a li

Full text of DOI:10.1021/jacs.9b02697

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Article
SiRA – A Silicon Rhodamine-Binding Aptamer
for Live-Cell Super-Resolution RNA Imaging
Regina Wirth, Peng Gao, Gerd Ulrich Nienhaus, Murat Sunbul, and Andres Jäschke
J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/jacs.9b02697 • Publication Date (Web): 15 Apr 2019
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SiRA – A Silicon Rhodamine-Binding Aptamer for Live-Cell Super-
Resolution RNA Imaging
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Regina Wirth†, Peng Gao‡,§, G. Ulrich Nienhaus‡,§,∥,⊥, Murat Sunbul*,† and Andres Jäschke*,†
† Institute of Pharmacy and Molecular Biotechnology (IPMB), Heidelberg University, 69120 Heidelberg, Germany
‡ Institute of Applied Physics (APH), Karlsruhe Institute of Technology (KIT), Wolfgang-Gaede-Str. 1, D-76131 Karlsruhe,
Germany
§ Institute of Nanotechnology (INT), Karlsruhe Institute of Technology (KIT), D-76344 Eggenstein-Leopoldshafen, Germany
∥ Institute of Toxicology and Genetics (ITG), Karlsruhe Institute of Technology (KIT), D-76344 Eggenstein-Leopoldshafen,
Germany
⊥ Department of Physics, University of Illinois at Urbana−Champaign, 1110 West Green Street, Urbana, Illinois 61801,
United States
ABSTRACT: Although genetically encoded light-up RNA aptamers have become promising tools for visualizing and tracking RNAs
in living cells, aptamer/ligand pairs that emit in the far-red and near-infrared (NIR) regions are still rare. In this work, we developed
a light-up RNA aptamer that binds silicon rhodamines (SiRs). SiRs are photostable, NIR-emitting fluorophores that change their
open-closed equilibrium between the non-colored spirolactone and the fluorescent zwitterion in response to their environment. This
property is responsible for their high cell permeability and fluorogenic behavior. Aptamers binding to SiR were in vitro selected from
a combinatorial RNA library. Sequencing, bioinformatic analysis, truncation and mutational studies revealed a 50-nucleotide minimal
aptamer, SiRA, which binds with nanomolar affinity to the target SiR. In addition to silicon rhodamines, SiRA binds structurally
related rhodamines and carborhodamines, making it a versatile tool spanning the far-red region of the spectrum. Photophysical
characterization showed that SiRA is remarkably resistant to photobleaching and constitutes the brightest far-red light-up aptamer
system known to date owing to its favorable features: a fluorescence quantum yield of 0.98 and an extinction coefficient of 86,000
M-1cm-1. Using the SiRA system, we visualized the expression of RNAs in bacteria in no-wash live-cell imaging experiments, and
also report stimulated emission depletion (STED) super-resolution microscopy images of aptamer-based, fluorescently labeled
mRNA in live cells. This work represents, to our knowledge, the first application of the popular SiR dyes, and of intramolecular
spirocyclization as a means of background reduction in the field of aptamer-based RNA imaging. We anticipate a high potential for
this novel RNA labeling tool to address biological questions.
dynamics and, concomitantly, to
a strong increase in
INTRODUCTION
fluorescence.8 This strategy is utilized by the 3,5-difluoro-4-
hydroxybenzylidene imidazolinone (DFHBI)-binding Spinach
and Broccoli aptamers, the thiazole orange-binding Mango
aptamers and their improved versions.13-18 Similarly, the
recently introduced DFHO-binding Corn aptamer allowed for
the visualization of RNA polymerase III activity.19 The
alternative strategy for RNA imaging uses bright, photostable
organic dyes with an appended quenching moiety that
suppresses the fluorescence in solution by contact quenching.8
Binding of the fluorophore or the quencher to the aptamer
disrupts the fluorophore-quencher interaction, leading to a
fluorescence increase.20 Our group previously reported the
promiscuous SRB-2 and DNB aptamers, which bind a variety
of fluorophores connected to the contact quencher
dinitroaniline.21-22 Analogously, Sato et al. have imaged
endogeneous RNAs using a BHQ (black hole quencher)-
binding aptamer combined with a BHQ-fluorophore probe.23-24
To gain insight into the complex life of RNA, the toolbox of
markers for RNA imaging has become quite diverse and is
continually expanding to adapt to state-of-the-art microscopy
techniques.1-7 Light-up RNA aptamers provide a particularly
attractive approach to label specific RNAs in live cells.8-10
These oligonucleotides bind small fluorogenic probes, typically
organic dyes, with high specificity and affinity, and are usually
generated in vitro using systematic evolution of ligands by
exponential enrichment (SELEX).11-12 The selected aptamer
sequence can be appended as a genetically encoded tag to the
RNA of interest and transcribed by the cellular machinery.
Binding of the probe to the aptamer triggers an increase in
fluorescence that can be visualized using fluorescence
microscopy. 2-3, 8
Several aptamer-ligand pairs spanning the VIS region have
been established for live-cell RNA imaging, using mainly two
strategies for fluorogenicity enhancement of the dye: restriction
of intramolecular movement, and disruption of a quencher-dye
conjugate. The former approach uses a chromophore that has a
low fluorescence quantum yield due to molecular motions in
solution. Aptamer binding leads to a suppression of the
Fluorescent dyes that can be excited and emit in the far-red and
near-infrared region are of particular interest for live cell
imaging because, in this spectral range, light is less phototoxic,
and cellular autofluorescence background is reduced.25-27
Moreover, such fluorophores are also compatible with laser
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wavelengths preferred in super-resolution microscopy.28-29
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Recent developments in the field such as the TO-3-binding
Mango, the malachite green-binding MG or the latest Riboglow
aptamer showed successful RNA visualization in the far-red
region.16,30-31 However, these light-up aptamers recognize dyes
that are less suited for live-cell imaging, as they are either
phototoxic, hardly cell-permeable, or not very bright.8,28,32-33
The DIR-pro system is the brightest far-red RNA aptamer
reported to date, but it suffers from low cell permeability and
therefore has only been used for imaging of a receptor on the
cell surface when fused to a receptor-binding aptamer.34 Thus,
there is a strong need for far-red and near-infrared aptamer
reporters which recognize bright, photostable and cell-
permeable dyes.
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The work presented herein focuses on the development of a
novel near-infrared light-up RNA aptamer that binds a cell-
permeable fluorophore, combined with a novel concept of
fluorogenicity in the field of aptamer-based RNA imaging,
namely environment-dependent spirocyclization. Silicon-
substituted xanthene dyes (silicon rhodamines, SiRs) have
become popular fluorophores for protein labeling since they are
photostable near-IR fluorophores compatible with standard
confocal and super-resolution microscopes.35-37 Among these,
3-carboxy SiRs exhibit an open-closed equilibrium between a
Figure 1. General RNA labeling strategy using a SiR-binding
aptamer. Top: environment-dependent intramolecular
spirocyclization of silicon rhodamines. Bottom: The fluorescence
increases upon binding of the aptamer to the SiR fluorophore,
allowing visualization of the target RNA using fluorescence
microscopy.
RESULTS AND DISCUSSION
Target Synthesis and Characterization. For the selection, we
aimed to generate 5-carboxytetramethylsiliconrhodamine (1)
(Figure 2A), since it carries two carboxyl functionalities: one
for the lactone-zwitterion equilibrium and a second one for
conjugation to a solid support, which is a necessity for the
SELEX protocol. We synthesized and characterized 1 using the
classical route38 via a 10-silaanthrone and a dioxazole-protected
arylhalide in four steps (Figure 2A). Next we prepared 2 by
coupling a 2,2’-(ethylenedioxy)bisethylenediamine linker to 1
(Figure 2A) and determined its photophysical properties.
non-colored spirolactone and
a fluorescent zwitterion,
depending on the environment (Figure 1). This property causes
high cell permeability in combination with a fluorogenic
behavior in solution and upon unspecific binding to cellular
components.38-39 These beneficial properties have been
exploited for imaging cellular proteins with unprecedented
resolution, and have inspired synthetic chemists to further vary
and optimize these probes.28, 36, 40-43 However, the application of
SiRs in the field of aptamer-based RNA imaging has not yet
been reported. We expected that the stabilizing environment of
an RNA aptamer combined with SiRs’ fluorogenicity could
induce sufficient fluorescence increase for RNA labeling
without the need of an additional quencher moiety (Figure 1).
Here, we report the selection, bioinformatic analysis,
mutational study, and characterization of a novel 50-nucleotide
(nt) RNA aptamer that binds a 3-carboxylated silicon
rhodamine with nanomolar affinities. Furthermore, using
confocal and super-resolution STED microscopy, we
demonstrate live-cell imaging experiments with this very
bright, near-IR labeling tool for RNA visualization. To the best
of our knowledge, this work represents the first example of
intramolecular spirocyclization as a means of background
reduction in the field of RNA imaging, and we envision a high
potential for this novel labeling strategy.
The extinction coefficient in aqueous buffer (27,000 M-1cm-1),
excitation and emission maxima (Exmax = 649 nm, Emmax
=
666 nm) as well as the fluorescence quantum yield (Φf = 0.51)
were all similar to values reported for SiRs in the literature.44
To analyze how the polarity of the medium influences the
lactone-zwitterion equilibrium, the absorbance of 1 and of its
tetramethylrhodamine and carborhodamine analogues, 3 and 4,
respectively, (for synthesis, see supporting information) was
measured in water/dioxane mixtures of varying composition.
As water is a polar solvent with a high dielectric constant (εr =
80.4), it promotes the formation of the zwitterionic form. By
contrast, dioxane is a nonpolar solvent with a low dielectric
constant (εr = 2.2), promoting the lactone form. We observed
that the dyes followed the expected trend: 1 and 2 (D0.5 ~ 60)
Figure 2. Synthesis and characterization of SiR 1 and 2. A) Synthetic scheme. B) Normalized absorbance of the different dyes versus the
dielectric constant of water/dioxane mixtures. 1 and 2 were compared to 5-carboxytetramethylrhodamine (3) and 5-carboxycarborhodamine
(4). The D0.5 value corresponds to the inter-section of interpolated graphs with Absorbance/Absorbancemax = 0.5.
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> carborhodamine (D0.5 = 20 - 40) > tetramethylrhodamine
increasingly harsher washing conditions (Figure 3B). To screen
(D0.5<10) (Figure 2B).28, 40 For example, 1 and 2 were mostly in
the eluted RNA pools for fluorescence enhancement,
dinitroaniline-quenched SiR probe 6 was prepared according to
our previously reported method (Figure 3A, for synthesis, see
supporting information).20, 22 We observed a gradual increase in
the fluorescence intensity of the probe when mixed with RNA
pool 7, 11 and 13 compared to the starting RNA library (Figure
S2), which indicates an enrichment of aptamers specifically
binding to the fluorophore.21 We reverse-transcribed selection
pool 14, PCR-amplified, cloned it into a plasmid and sequenced
individual colonies by Sanger sequencing (Table S2).
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the nonfluorescent spirolactone form in water/dioxane mixtures
with
a
dielectric constant of less than 40, where
tetramethylrhodamine favored predominantly the fluorescent
zwitterionic form, and carborhodamine existed as a mixture of
spirolactone and zwitterion forms. Since SiR 2 met all
requirements for aptamer selection, we immobilized it on
sepharose beads and used those in the SELEX protocol.
Selection. The library used for aptamer selection had a diversity
of ~2×1015 and a standard length of 103 nt, where each
individual molecule carried three key features (Figure 3 A and
Table S1): (1) a 12-nt stable stem-loop, which has been shown
to increase the probability of finding high-affinity aptamers,45
(2) two fixed primer binding sites for reverse transcription and
amplification, and (3) two 26-nt random regions for diversity.
After transcribing the initial DNA library, the RNA was mixed
with the Sepharose beads functionalized with 2 and subjected
to the iterative SELEX protocol, in which the output of one
round serves as input for the next one. The protocol included
bead binding and washing, elution of bound RNAs,
amplification and, furthermore, involved counter-selection
against sepharose binders (Figure S1). Figure 3B shows an
exponential increase of the eluted RNA fraction over 7 rounds
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Bioinformatic Analysis and Motif Investigation. In order to
identify a common binding motif, phylogenetic analysis of the
sequences was carried out, revealing a total of 30 different
families (Figure S3).46 Next, we screened at least one sequence
in each clustered family in the fluorescence turn-on assay
(Figure S4). Out of 42 sequences, 21 showed no significant
turn-on upon mixing with probe 6 and were excluded from
further analysis. Sequence alignment of all fluorescence-
activating aptamers revealed a motif conserved in 70% of the
sequences (Figure S5 and Figure 4A).
Figure 4. Characterization and truncation of SiR-binding aptamers.
A) Predicted three-way junction motif derived from the SELEX
data. B) Predicted secondary structure of the 50-nt mutant M1
(SiRA). The conserved motif, which was found in 70 percent of the
active clones, is highlighted in color. C) KD curves of SiRA and
full-length C02. KD (C02): 490 ± 140 nM, KD (M1): 430 ± 70 nM.
50 nM 2 was incubated with different concentrations of RNA.
Figure 3. SELEX of SiR-binding aptamers. A) Design of the
partially structured RNA library and structures of SiR-biotin (5)
and contact-quenched SiR-dinitroaniline (6). B) Progress of the in
vitro selection was monitored by the percentage of RNA eluted
from the beads functionalized with either ligand 2 or 5.
The overall shape of the motif is a three-helix junction. H1 is
attached to the primer A and B binding site (blue) and evolved
together with H2 from the random portions of the library,
whereas helix H3 (yellow) is the one incorporated into the pool
by design (Figure 3A). While the length of H3 is rather constant,
H1 and H2 are variable (Figure 4A). In some sequences, H1,
H2, and H3 are connected directly to each other, while in other
cases, there are interspersed linker nucleotides. H3 is then
connected to H1 via a formally single-stranded loop (L1) that
harbors a highly conserved UAUC tetranucleotide (green)
sequence, which is located 2-15 nt away from H3. The loop that
of selection, which indicates enrichment of target-binding
RNAs.22 To increase the selection pressure, 2 was conjugated to
a biotin moiety to yield ligand 5, which can be immobilized on
streptavidin-agarose beads with a controlled stoichiometry and
selectively cleaved from the beads via S-S bond reduction upon
DTT addition (Figure 3A). Using ligand 5, 7 more rounds were
carried out using decreasing dye concentrations and
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closes H2 and L2 (UGAA, pink) is also highly conserved in combinations (M13, M14, M15, M16). Although the
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sequence, including the two closing base pairs of H2 (2 A-U
pairs, pink). To find the smallest fluorescence-enhancing
aptamer, we analyzed sequence C02 that contained all features
in the most condensed form (Figure S6), and also showed the
highest fluorescence enhancement in the turn-on assay (Figure
S4). We prepared a short 50-nt mutant (M1, SiRA), deleting the
3’- and 5’-extensions and introducing two guanosines at the 5’-
end for efficient in vitro transcription by T7 RNA polymerase
(Figure 4B). We compared the binding affinity of full-length
aptamer C02 and shortened SiRA aptamer for the selection
target 2 by titration of fixed amounts of 2 with the RNA (Figure
4C). Dissociation constants were found to be almost identical
and in the nanomolar range (430 ± 70 nM for SiRA and 490 ±
140 nM for the full-length C02 aptamer), and are thus
comparable to the well-established Spinach system. The
observation that shortening of the aptamer from 103 to 50 nt did
not lead to any loss of its binding affinity strongly supports the
view that the identified common three-way junction is the SiR
binding motif.
fluorescence increased slightly (M15, M16), the measurement
of the affinity of M15 (exchange to a stronger G-C bond)
towards 2 showed an increase in KD to ~2000 nM (Figure S10).
The exchange of the same G-U pair to an A-U pair increased
the KD to a high micromolar level. Insertion of a single
nucleotide (M17) at the AU of the closing stem H1 also
completely disrupted the binding, thus indicating that these
positions are crucial for proper aptamer folding. Taken all
bioinformatic and fluorescence enhancement data together, we
have successfully identified a three-way junction 50-nt RNA
aptamer (SiRA) that binds with nanomolar affinities to probe 2.
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Spectroscopic Characterization of the SiRA System. Figure
6A shows the absorption and emission spectra of probe 2 and
the [SiRA*2] complex. The absorbance of [SiRA*2] is roughly
threefold higher than that of unbound 2, indicating aptamer
binding to and shifting the equilibirium towards the zwitterionic
form of 2. We observed a ~7-fold fluorescence turn-on due to a
combination of two phenomena: the increase in extinction
coefficient of the [SiRA*2] complex, and the increase in the
fluorescence quantum yield to almost 1 (Table 1, Table S3).
To assess the influence of individual loops, helices and
nucleotides in the proposed motif, we generated a total of 16
mutants and assayed them for fluorescence enhancement using
contact-quenched probe 6 (Figure 5, Figure S7-9).
Table 1. Photophysical properties of silicon rhodamine 2
and [SiRA*2].
Brightnessc
b
λex
λem
ε
(nm)a
(nm)a
(M-1cm-1)a
27000
86000
2
649
666
662
0.51 13770
0.98 84280
[SiRA*2] 649
a
Determined in 20 mM HEPES (pH 7.4.), 5 mM MgCl2 and 125
mM KCl. b Determined in 20 mM HEPES (pH 7.4.), 5 mM MgCl2
and 125 mM KCl buffer using Oxazine 1 as standard (For details
see supporting information). c Brightness =   .
As a control, we incubated 2 with total RNA and observed a
slight decrease (~15%) in fluorescence, which has been
reported for non-specific RNA-dye interactions.21-22 The
significant increase in fluorescence quantum yield upon
aptamer binding is presumably caused by restricted movements
of the fluorophore and the hydrophobicity of the binding pocket
in the RNA. Thus, the [SiRA*2] system is currently the
brightest and most red-shifted system; it is 2.6 fold brighter than
eGFP and ~1.5 fold brighter than DIR-pro (Table S4).34, 49
Figure 5. Alignment of the SiRA mutants. The relative
fluorescence in the turn-on screening is given in comparison to M1
(SiRA). The mutations are highlighted in red, the identified
common motif is highlighted in color according to Figure 4.
The main loop L1 was found to be crucial for keeping the
tertiary structure competent for SiR binding. A single mutation
in the conserved UAUC sequence (M2) as well as a further
exchange of a UU sequence with CC (M3) abolished
fluorophore binding. Subsequently, we focused on the
tetraloops L2 and L3. Exchange of the UUCG with a different
stable GAAA tetraloop in L3 (M4) disrupted the complex,47
indicating that these nucleotides do not just stabilize stem H3,
but make important tertiary interactions. Next, we probed
potential “kissing interactions” between loops L2 (UGAA) and
L3 (UUCG) by exchanging up to three nucleotides on both
loops (M5, M6, M7 and M8), finding no support for this
hypothesis.48 In addition, further truncations of M1 by 6-nt
deletions from both 5´- and 3´- ends (M9) disrupted the
fluorophore RNA complex. We also investigated the influence
of the helix lengths in H1, H2 and H3. Having a 5 or 6 nt long
H1 stem (M10) did not make any difference and elimination of
one base-pair in H3 only slightly decreased the activity (M11).
On the other hand, the fluorescence decreased strongly upon
shortening H2 by two base-pairs (M12). This indicates that the
H2 hairpin length is important for folding. Next we tried to fix
Next we investigated the influence of temperature on the
[SiRA*2] system (Figure 6B). The fluorescence intensity of
[SiRA*2] decreased between 20 and 70 °C. At 37 °C, the
aptamer fluorophore complex still retains 78 ± 0.6% of its
fluorescence, which should render it well suitable for live-cell
imaging at this commonly used temperature and is comparable
to other aptamers.21, 50 At 47 °C, the fluorescence decreased to
50 ± 0.4% of its initial value. Since aptamer folding is
dependent on Mg2+ ion concentration, we also tested the
fluorescence of [SiRA*2] at different Mg2+ concentrations
(Figure 6C). The SiRA system retains 70-90% of its
fluorescence in the concentration range between 0.25-1 mM,
and the fluorescence slightly increases between 1-10 mM,
reaching its maximum value at 10 mM Mg2+. This experiment
shows that intracellular free Mg2+ in bacteria or mammalian
cells, regulated by transporters and generally maintained in the
range of 0.2-1 mM,51 is sufficient for the SiRA system to fold
correctly and should allow live-cell RNA imaging with SiRA.
a
G-U base-pairing “error” in helix H2 by different
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Figure 6. Characterization of the [SiRA*2] complex. A) Normalized absorption and emission spectra of free SiR 2 and the [SiRA*2]
complex. Emission of 2 with total RNA is also shown (cRNA= 20 µM , c2 = 1 µM). B) Temperature dependence of the [SiRA*2] complex
(cSiRA = 20 µM , c2 = 20 µM) determined in 20 mM HEPES (pH 7.4), 5 mM MgCl2 and 125 mM KCl. C) Magnesium dependence of
[SiRA*2] determined in 20 mM HEPES (pH 7.4.) and 125 mM KCl, (cSiRA = 20 µM , c2 = 20 µM).
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Promiscuity of the SiRA Aptamer. Since SiRs are structurally
very close to carborhodamines, which have recently gained
S11). Removal of the 2,2’-(ethylenedioxy)bisethylenediamine
linker from SiR 2 leads to a five-fold increase in KD ([SiRA*1]).
This indicates specific interactions between SiRA and either the
amide bond or the amino-triethylene glycol linker. Such strong
interactions between an aptamer and the whole ligand used for
solid phase support attachment have been reported.54
interest due to enhanced photostability and intermediate
52-53
spirocyclization behavior,28,
we compared the binding
properties
between
the
structurally
related
tetramethylrhodamine (3), carborhodamine (4) and silicon
rhodamine (1) analogues and the SiRA aptamer in vitro (Figure
7A).
Table 2. Properties of the different SiRA complexes used in
this study.
λex
λem
KD (nM)a
Turn-onb
(nm)a
(nm)a
[SiRA* 2]
[SiRA* 1]
[SiRA* 4]
[SiRA* 3]
649
649
613
559
650
662
660
627
575
662
430 ± 70
2600 ± 220
900 ± 130
3400 ± 320
1000 ± 230
7
6
3
1.5
17
[SiRA* 6]
a
measured in 20 mM HEPES (pH 7.4), 5 mM MgCl2 and 125
mM KCl at 25°C. b Fluorescence turn-on was determined from
the KD curve (max/min).
In addition, the free carboxylic acid in 1 might cause a repulsive
interaction with the negatively charged RNA backbone.55
Interestingly, carborhodamine 4 binds to SiRA approximately
threefold more strongly than 1, while the affinity for rhodamine
3 decreased slightly (1.3-fold) with respect to 1. These findings
illustrate that the bridging atoms (O, C(CH3)2, Si(CH3)2) of the
xanthene system are recognized by the aptamer, and that small
differences in the structure translate into significant changes in
the binding affinity. To investigate these effects,
crystallographic analysis could be of great advantage and will
be addressed in the future. Overall, the fluorescence turn-on
values of the SiRA-dye complexes were in agreement with the
Figure 7. Promiscuity of the SiRA aptamer. A) Structure of the
dyes binding to SiRA: 5-carboxytetramethylrhodamine (3), 5-
carboxytetramethylcarborhodamine
(4),
5-carboxysilicon-
previously
tetramethylrhodamine,
reported
open-closed
carborhodamines
equilibria
and
of
silicon
rhodamine (1). B) Normalized emission spectra of the different
[SiRA*dye] complexes upon excitation at 590 nm (1), 510 nm (3),
and 557 nm (4).
rhodamines (Table 2, Figure S11).28, 52-53 3, which is preferably
in the zwitterionic form, shows only a 1.5-fold increase,
followed by carborhodamine 4 with a threefold increase. SiRs
1 and 2, which have a higher tendency to be in the lactone form
when unbound, show the highest increase (six- to sevenfold).
One approach to quickly increase the fluorogenicity is the
combination of our previously reported contact-mediated
Figure 7B shows the emission spectra of all three rhodamines
in complex with the SiRA aptamer, spanning the far-red region
and demonstrating SiRA’s promiscuity. The equilibrium
dissociation constants, KD, vary by a factor of ~7 and give
insight into the determinants of dye binding (Table 2, Figure
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quenching with SiRA’s novel fluorogenicity concept. Initial in Repeats of the SiRA aptamer were used in order to enhance the
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vitro measurements with the contact-quenched SiR probe 6
showed a 17-fold increase in fluorescence (Table 2). However,
the dissociation constant increased twofold to a low micromolar
range, likely due to the extra energy required for the disruption
of the fluorophore-quencher complex.
signal. Figure 9 (488 nm channel) shows similar expression
levels of the GFP protein in the control and in the SiRA
construct, indicating no influence of the SiRA aptamer tag on
protein expression, while bright, localized fluorescence
emission upon 640-nm excitation was only detected in the SiRA
construct (Fig. 9B). We predominantly observed accumulation
of mRNA at the poles of E. coli (Figure 9C), as reported
previously.56,57 The average fluorescence intensity at the poles
of the SiRA-expressing bacteria was found to be ~34 times
higher than that of the control bacteria expressing only GFP
(Figure S12). It should be noted that this experiment involved
neither washing nor a quenching moiety.
No-wash, No-quencher RNA Visualization in Live Bacteria.
Since not only sufficient fluorogenicity but also low
dissociation constants are key requirements for successful
aptamer-based RNA imaging, we chose the [SiRA*2] system
for proof-of-principle imaging experiments. In addition, a free
amine handle on the silicon rhodamine that allows for further
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functionalization can be seen as
a benefit for future
applications, e.g., investigating RNA-RNA or RNA-protein
interactions. We prepared a pET28 expression vector carrying
one copy of the SiRA aptamer embedded in a tRNA scaffold
under the control of the T7 promoter and terminator. Upon
addition of IPTG, SiRA was expressed and visualized by
addition of probe 2. Figure 8 shows in E. coli cells labeled with
[SiRA*2] in comparison to control bacteria. The bright signal
in the red channel indicates proper folding of the SiRA aptamer
in live cells. This experiment shows the use of a photostable,
far-red, bright fluorophore without the need of an additional
quenching moiety in a no-wash live-cell RNA labeling
experiment.
Figure 9. Imaging of GFP mRNA in live E. coli bacteria with a
confocal microscope. A) Expression scheme of GFP mRNA, lacO:
lac operon, rbs: ribosomal binding site, ROI: RNA of interest, P:
promoter, T: terminator. B) No-wash, live-cell imaging of GFP
mRNA in live E. coli cells. The E. coli were transformed with either
a plasmid carrying GFP (control) or GFP-SiRA5. GFP expression
was induced by addition of IPTG (1 mM). The cells were incubated
for 5 min with 500 nM of 2 and then directly imaged at 37 °C using
a confocal microscope. Scale bar 5 µm. C) Zoom of the mRNA
expression images depicted in Figure 9B. Scale bar 5 µm.
To analyze the dynamic range of the [SiRA*2] system, we
imaged fixed cells as a function of dye concentration between
50 and 1000 nM (Figure S13). We chose fixed cells because the
RNA concentration remains constant over time. High
fluorescence intensity in SiRA-expressing bacteria indicated
full function of [SiRA*2] after paraformaldehyde fixation. The
ideal concentration of 2 was determined as 500 nM, as lowering
2 below the KD decreased the detected amount of mRNA.
Increasing 2 to 1 µM caused higher background fluorescence in
the control bacteria that did not express SiRA.
Figure 8. No-wash, live-cell imaging of tRNA expression in E.
coli. A) SiRA-RNA vs. tRNA expression scheme, P: promoter, T:
terminator B) Brightfield and fluorescence images with 640-nm
excitation. Cells were transformed with either a pET28 vector
expressing the SiRA aptamer in a tRNA scaffold or only the tRNA
construct as a negative control. The transcription of the aptamer
was induced by addition of IPTG (1 mM). The cells were incubated
for 10 min with 500 nM of 2 and then directly imaged at 37 °C
using a wide-field microscope.
To show labeling of less stable RNAs, we prepared a plasmid
for expression of GFP mRNA carrying 5 repeats of the SiRA
aptamer inserted into the 3´ UTR without a stabilizing tRNA
scaffold, which we denote GFP-SiRA5 mRNA in the following.
Next, we modulated the expression levels of GFP mRNA (Fig.
S14) by varying the amount of added IPTG (1000 - 0 µM),
which controls the transcription upon lac repressor binding. The
fluorescence decreased in both 488 nm and 640 nm channels
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upon IPTG reduction. Below 25 µM IPTG, specific GFP-SiRA5 the fluorescence emission over time (Figure 10E). Indeed, the
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mRNA fluorescence was not detectable (Fig. S15). mRNA
levels were quantified by RT-qPCR (Table S5), revealing a
sixfold reduction of RNA expression at 25 µM IPTG with
respect to 1 mM IPTG. According to the RT-qPCR data, each
pole of a bacterium contained on average between 300-700
copies of GFP-SiRA5 mRNA at 25 µM IPTG. The visualization
of lower expression levels with this setup will therefore require
constructs with higher repeat numbers (e.g., SiRA24), as is
commonly done in other RNA imaging systems.1,57-58
[SiRA*2] complex was much more photostable than Atto 647N
in solution.
9
Super-resolution Microscopy of mRNA in Live Bacteria.
Fluorogenicity, brightness, photostability and near-IR emission
of SiR dyes make them ideal fluorophores for super-resolution
microscopy (SRM). They have already been widely used for
super-resolution imaging of proteins,38-39, 59 and so we tested
[SiRA*2] for RNA visualization using SRM. Figure 10 shows
stimulated emission depletion (STED) microscopy images of
live bacteria expressing GFP-SiRA5 mRNA.60 The STED
images (Figure 10A, insets: time sequence taken every 31.5 s)
show much sharper features than standard confocal images
(Figure 10B) due to the enhanced resolution of ~40 nm (Figure
S16, for experimental details, see Supporting Information). We
note that, whereas SRM of proteins is well established,61-63
applications to RNA are extremely rare and usually limited to
fluorescence in situ hybridization (FISH) based methods.64-65
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During STED imaging of GFP-SiRA5 mRNA, excellent
photostability of the [SiRA*2] system was observed. Under
settings where a common STED standard (fluorescent dark-red
beads, see Supporting Information for details) was
photobleached to ~10%, [SiRA*2] fluorescence in fixed
bacteria decreased to only ~60% of its inital value (Figure 10C).
To characterize the response of the [SiRA*2] system to light
irradiation, we repetitively scanned fixed bacteria labeled with
GFP-SiRA5 mRNA using focused 640 nm laser light of 8 or 30
µW power and measured their fluorescence emission as a
function of time (Figure 10D). The quantitative analysis, which
is described in detail in the Supporting Information, yields a
bleaching rate coefficient, kl, of the [SiRA*2] complex of 0.67
± 0.04 × 10-3 s-1 µW-1, indicating that it would take ~103 s to
bleach half of the aptamer-bound SiR molecules under 1 µW
irradiation. Because the fluorescence can be restored by
exchange of photobleached dyes by emissive ones from the
solution, the analysis also yields the dissociation rate
coefficient, kd, of [SiRA*2] as 2.8 ± 0.2 × 10-3 s-1. An essentially
identical value was obtained by a fluorescence recovery after
photobleaching experiment in fixed bacteria expressing GFP-
SiRA5 mRNA (Fig. S16). The off-rate of the [SiRA*2] system
is 2.9 and 6.4-fold lower than that of the reported Corn-DFHO
Figure 10. STED imaging of live E. coli bacteria expressing GFP-
SiRA5 mRNA and photostability analysis of [SiRA*2]. The
bacteria were transformed with a plasmid carrying GFP-SiRA5;
GFP expression was induced by addition of IPTG (1 mM). The
cells were incubated for 5 min with 500 nM of 2 and then directly
imaged at 37 °C. A) Bottom: Time series of STED images taken in
31.5 s intervals for a total of 126 s; top: magnified image at 126 s;
image parameters: 200  200 pixels, 30 µs per pixel, 640 nm
excitation (5 µW) and 779 nm depletion (150 mW). B) Regular
confocal image of GFP-SiRA5 expressing bacteria, taken directly
after the STED time series. C) Fluorescence decay of the [SiRA*2]
aptamer in a fixed E. coli cell under continuous STED image
scanning, using 640 nm excitation (5 µW) and 779 nm depletion
(70 mW). For comparison, we show the behavior of dark-red beads
under the same conditions. D) Normalized fluorescence decay of
the [SiRA*2] aptamer in E. coli cells upon repetitive scanning (200
 200 pixels, 30 µs per pixel) with a 640 nm laser at 8 and 30 µW
power. E) Normalized fluorescence of [SiRA*2] and Atto 647N
and Broccoli-DFHBI-1T systems, respectively.19 Since KD
=
kd/ka, the association rate constant ka of [SiRA*2] is 6.5 ± 1.1 ×
103 M-1 s-1 and thus 3.5- and 8-fold lower than in DFHO and
Broccoli-DFHBI-1T, respectively.19
Finally, we compared the photostability of Atto 647N, which is
a very photostable dye commonly used in STED,66-69 and the
[SiRA*2] complex in solution. Since the fluorescence decay in
the imaging and time-dependent irradiation experiments is
affected by both photobleaching and dye-aptamer exchange in
the presence of excess dye, we prepared a sample with an
excess of SiRA aptamer (15 nM of dye and 10 µM of the
aptamer), so that no fluorescence recovery occurs after
photobleaching. We then illuminated both [SiRA*2] and Atto
647N solutions with a 645 nm LED lamp for 600 s and recorded
upon photobleaching with
a high-power 645 nm LED;
concentrations of Atto 647N and 2 are both 15 nM.
Summary and Outlook. In summary, we have presented the
selection, characterization and application of
a silicon
rhodamine-binding RNA aptamer – SiRA. Photophysical
characterization showed a significant fluorescence turn-on upon
dye-aptamer binding, generating an extremely photostable and
the brightest far-red light-up aptamer system reported to date.
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3. Xia, Y.; Zhang, R.; Wang, Z.; Tian, J.; Chen, X., Recent advances
In a proof-of-principle study, we employed the SiRA system to
visualize the expression of mRNA in bacteria in no-wash,
quencher-free, live-cell imaging experiments, thereby
introducing intramolecular spirocyclization as a novel concept
of fluorogenicity to the field of aptamer based RNA imaging.
In addition, we have successfully applied the SiRA system in
the first light-up aptamer application of live-cell SRM of
mRNAs. We have limited ourselves to prokaryotic systems in
this work, but the application of SiRA to mammalian cells is
currently under investigation. To image RNAs of low
abundance and to increase the sensitivity of the system, a large
number of tandem repeats of SiRA can be genetically fused to
the RNA of interest. Further selection, mutagenesis and
engineering might yield even brighter, less ion-sensitive and
more thermostable aptamers with even lower equilibrium
dissociation constants.70 Another optimization goal for future
studies would also be increasing the relatively small Stokes
shift. We observed that the emission maxima are slightly blue
shifted upon binding of SiRA to all substrates (Table S3).
Although it will be challenging to generate high-affinity
aptamers for dyes with a high degree of spirocyclization, we
envision that a combination of our new labeling concept with
the recently reported Janelia dye palette or the spontaneously
blinking hydroxymethyl-SiR could further increase the
fluorescence turn-on.41, 71 Therefore, we expect that SiRA will
become a valuable addition to the toolbox of RNA imaging and
we are excited about further evolving this novel labeling
concept towards its application in eukaryotic systems.
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ASSOCIATED CONTENT
The Supporting Information is available free of charge on the ACS
Publications website.
Experimental details and supporting figures (PDF). “This material
is available free of charge via the Internet at http://pubs.acs.org.”
AUTHOR INFORMATION
Corresponding Author
* jaeschke@uni-heidelberg.de
* msunbul@uni-heidelberg.de
Funding Sources
A.J. was supported by the Deutsche Forschungsgemeinschaft
(Grant # Ja794/11-1) and G.U.N. by the Helmholtz Association
(Program Science and Technology of Nanosystems).
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENT
We thank Thorge Reiber, Katharina Peters and Fanny Sanchez for
experimental support, the Nikon Imaging Center, Heidelberg for
granting access to their facilities and Dr. Christian Ackermann for
technical advice in fluorescence microscopy.
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