Life times of metastable states guide regulatory signaling in transcriptional riboswitches

ABSTRACT Transcriptional riboswitches modulate downstream gene expression by a tight coupling of ligand-dependent RNA folding kinetics with the rate of transcription. RNA folding pathways

leading to functional ON and OFF regulation involve the formation of metastable states within well-defined sequence intervals during transcription. The kinetic requirements for the formation

and preservation of these metastable states in the context of transcription remain unresolved. Here, we reversibly trap the previously defined regulatory relevant metastable intermediate of

the _Mesoplasma florum_ 2′-deoxyguanosine (2′dG)-sensing riboswitch using a photocaging-ligation approach, and monitor folding to its native state by real-time NMR in both presence and

absence of ligand. We further determine transcription rates for two different bacterial RNA polymerases. Our results reveal that the riboswitch functions only at transcription rates typical

for bacterial polymerases (10–50 nt s−1) and that gene expression is modulated by 40–50% only, while subtle differences in folding rates guide population ratios within the structural

ensemble to a specific regulatory outcome. SIMILAR CONTENT BEING VIEWED BY OTHERS REAL-TIME MONITORING OF SINGLE ZTP RIBOSWITCHES REVEALS A COMPLEX AND KINETICALLY CONTROLLED DECISION

LANDSCAPE Article Open access 10 September 2020 A NASCENT RIBOSWITCH HELIX ORCHESTRATES ROBUST TRANSCRIPTIONAL REGULATION THROUGH SIGNAL INTEGRATION Article Open access 10 May 2024 RNA

FOLDING KINETICS CONTROL RIBOSWITCH SENSITIVITY IN VIVO Article Open access 22 January 2025 INTRODUCTION Riboswitches selectively bind metabolites to control the transcription or translation

of metabolite-associated genes. In general, riboswitch function is described as a ligand-dependent switch between two mutually exclusive conformations to either permit or repress gene

expression1,2. For riboswitches that regulate gene expression on the level of translation, reversible conformational switching allows RNAs to adopt distinct ligand-dependent functional folds

at equilibrium during multiple rounds of translation. As a consequence, RNA synthesis and the cellular response to changes in metabolite concentration are separated in time. The functional

relevant switch underlying translational riboswitches modulate ribosome access to the SD sequence3,4,5,6, but also more complex regulation mechanisms have been discovered7. In contrast,

control of gene expression by premature termination of transcription inevitably requires temporal coupling of RNA synthesis and conformational switching. Riboswitches utilize rho-independent

transcriptional termination8,9. Under these conditions, the nascent transcript can either adopt a sequence-conserved terminator conformation that disrupts the polymerase complex or form one

or several alternative, so-called antiterminator conformations that avoid termination. The metabolite concentration EC50, at which riboswitches operate, often exceeds the KD values for

ligand binding by at least one order of magnitude3,10,11,12,13,14. This observation indicates that fast folding of functional states during well-defined transcription intervals is important

for riboswitch function. In a previous study, we investigated the landscape of the thermodynamically stable states adopted in either the presence or absence of metabolite for mRNA

transcripts of increasing chain length15. The ligand-dependent conformational rearrangement underlying riboswitch function typically involves four distinct sequence segments P, A, T, and H

as depicted in Fig. 1. In the 2′dG riboswitch, an mRNA containing 80 nucleotides is the first transcript that can form the interaction between strands P and A. Ligand binding can occur

directly after the aptamer domain is transcribed. Interestingly, ligand can also bind to the growing mRNA up to nucleotides 80–113 nt, but the stability of these mRNA-ligand complexes and

thus the population of ligand-bound states continuously decrease during the addition of nts 90–113 from >90% to 70%. As strand T is transcribed, the PA interaction becomes metastable

(PA–T) in both the ligand-bound and ligand-free form. In the equilibrium model, folding from PA–T to the antiterminator conformation PAT concomitantly occurs during transcription of

nucleotides 113 to 137. Previous studies assume ligand binding within the first transcription interval (nts 80–110) to stabilize the PA interaction sufficiently to inhibit an interaction

with strand T and to ensure OFF function, but it remains uncertain how the PAT state is selected from only one metastable state in the context of transcription. PAT states adopted within the

second transcription interval (in green) become metastable as strand H is transcribed (PAT–H) during synthesis of nts 139 to 144 in both, absence or presence of ligand. Within this short

period during transcription, the terminator conformation PA–TH represents the most stable state. It follows that deciphering the change of kinetics of antiterminator formation (PAT)

depending on the absence and presence of ligand represents the most important information regarding regulation. According to the equilibrium model, the effects of ligand binding on the

overall fold adopted at a specific transcript length is insignificant. Rather, formation of metastable intermediate states as important navigators of the regulatory outcome suggests a

correlation between the life times of these states and transcription rates to be critical for riboswitch function. Here, we unravel these kinetic requirements to achieve and to maintain a

ligand-dependent reversal of population ratios of functional OFF and ON states during transcription. On the example of the central metastable PA–T conformation of the 2′dG riboswitch, we

demonstrate that ligand binding does not induce major changes in equilibration kinetics of the PA–T state, but slows refolding to allow the polymerase to bypass the metastable transcription

barrier for the majority of nascent transcripts. RESULTS REVERSIBLE TRAPPING OF THE METASTABLE PA–T CONFORMATION To monitor antiterminator (PAT) folding, we trapped the PA conformation of

the 121 mer transcriptional intermediate by incorporating photolabile protection groups in strand T. These protection groups prevent antiterminator helix PT formation and allow monitoring of

folding from PA to PT after photolysis by light-induced real-time NMR as reported previously16,17,18. The 121 nt transcriptional intermediate forms a complete antiterminator helix PT at

equilibrium. While refolding from PA–T to PAT is initiated at nucleotide 113, segment 113–121 only contains four weak base pairs within helix PT, and, therefore, refolding kinetics from PA–T

to PAT are most accurately described at length 121 on average. Photocaged dGsw121 (dGsw121,caged) was prepared by enzymatic splinted ligation and contained three NPE protection groups at

the C4 atoms in the nucleobases of Uracil residues 108, 110, and 112 (Fig. 2a, Supplementary Fig. 1). In addition to steric hindrance, these NPE protection groups introduce a tautomeric

shift at the Watson–Crick base pairing site to prevent base pairing. The RNA fragment comprising nucleotides 1–85 was transcribed in 15N-G/U isotope labeled form. The NPE-protected smaller

fragment comprising nucleotides 86–121 was chemically synthesized (Supplementary Fig. 2, Table 1). We analyzed dGsw121,caged by NMR-spectroscopy to verify that trapping the PA fold in the

caged construct was successful. Figure 2b shows an overlay of NMR spectra before and after photolysis, while in the 15N-correlation spectra only the 15N-labeled fragment (nts 0–85) is

detectable. In dGsw121,caged, the conformation of the aptamer domain (PA–T) is maintained as evidenced by characteristic imino proton reporter signals, in particular U17 and U1819.

Irradiation at 355 nm releases native dGsw121 and subsequently induces formation not of a single, but of two interconverting antiterminator conformations (PAT(I)) and (PAT(M)), identified by

helix PT reporter signals G12, U108, U109, and G25I. In the presence of ligand, the antiterminator conformations PAT folds only to ~80%, in agreement with the static experiments15. In the

ligand-free form, chemical shifts of helices P2 and P3 are identical in either the aptamer PA or antiterminator PAT conformations, while those signals can be distinguished in the

ligand-bound form due tertiary interactions induced by ligand binding. LIFE TIMES OF THE METASTABLE PA–T STATE BY REAL-TIME NMR We obtained kinetic traces by real-time NMR at the

physiological temperature of the bacterium _M_. _florum_ (25 °C). Rates for individual base pairs were determined (Supplementary Fig. 3), utilizing F-statistics to check for statistical

significance (Supplementary Table 2). Rapid formation of PAT in the absence of ligand required averaging of transients obtained from individual imino protons to yield a final rate for helix

formation with improved accuracy (Fig. 3a). We can confirm the validity of this approach by the larger number of kinetic traces with improved S/N obtained for the ligand-bound form (Fig. 

3b), where the average rate for helix P2/P3 formation accurately reflects rates of individual bases. In the absence of ligand, PAT(M) folds at a rate of ~1.3 s−1. The second conformation

PAT(I) could be monitored by the characteristic and isolated signal G25I, and folds at 10-fold slower rates of 7*10−2 s−1. In the ligand-bound form, the difference in chemical shift of

helices P2/P3 in PA–T and PAT allows monitoring of aptamer dissociation in addition to PT formation. The ligand-bound PA conformation folds to PAT(M) at rates of ~9*10−2 s−1 with a second

slower rate in the order of ~1*10−2 s−1, which we assign to re-equilibration to PAT(M) after formation of PAT(I). The analysis of the kinetics reveals three relevant functional pathways as

depicted in Fig. 1. Folding of PAT(M) in the absence of ligand (pathway 2) is on the same timescale (1.3 s−1) as bacterial polymerases require to synthesize the subsequent 24 nucleotides

(0.4–3.7 s−1)20,21,22,23. The synchronization of transcription with folding rates that are unimolecular and thus concentration independent and sufficiently long life-time of the metastable

state ensures ON state function. In contrast, ligand binding slows down folding from the PA conformation to the PAT(M) conformation to a rate of ~0.1 s−1, and, thus, below transcription

rates (pathway 3). Hence, in the presence of ligand, antiterminator formation for transcript length for which it represents the stable fold (113–137 nts) is too slow, and, thus, OFF state

function is triggered. Further, equilibration between PAT(M) and PAT(I), occurs on the same timescale as antiterminator formation in the presence of ligand, and, therefore, cannot compete

with transcription rates. Therefore, our kinetic experiments reveal that only one of the two antiterminator conformations (PAT(M)) represents the regulatory decisive ON state, and, thus,

demonstrate a kinetic selection of only a single functionally relevant conformation in the structural ensemble. LIGAND BINDING KINETICS OFF state function depends on ligand binding kinetics

and subsequent RNA folding. We further determined the kinetics for transcripts of different chain length by stopped-flow spectroscopy with a surrogate fluorescence ligand 2-aminopurine

2′-deoxyriboside (2′dAP) to investigate whether the ligand association rates (kon) remain constant within the relevant transcription interval. To facilitate binding of 2′dAP, the ligand

selectivity was shifted from G to A by mutating the Watson–Crick ligand recognition site C74 to U74 (dGswC74U), a common approach applied for purine riboswitches (Fig. 4a)24,25. Homogeneous

binding of 2′dAP to dGswC74U was verified by NMR and kon rates were obtained for 9 different transcriptional intermediates (Supplementary Fig. 4 and Table 3). The results show that ligand

binding kinetics are virtually identical for constructs that adopt a stable PA fold based on static measurements (Fig. 4b, transcript lengths 80–113 nt). Truncated P1 constructs as well as

antiterminator constructs exhibit a reduced kon rate by a factor of 5 and 2, respectively. Since KD values of 2′dAP binding to dGswC74U (60 µM) exceeds the KD of 2′dG binding to dGsw (250 

nM), the determined koff for 2′dAP of 0.35 s−1 provides an upper limit for koff of 2′dG. Analysis of ligand binding by stopped-flow spectroscopy shows that ligand binding kinetics remains

constant for stably formed aptamer domains regardless of transcript length. Due to decline in the ligand-bound equilibrium population between transcript length 90 and 110, the amount of

structures accumulating within pathway 3 depends on cellular ligand concentration and corresponding kinetics of ligand association. Therefore, at saturating ligand concentration, the bound

state (pathway 3) will be populated in a range of 70–100% by the time the polymerase has reached nucleotide 113. Equilibration can take place during transcription of nts 90–110, transcribing

at rates of 0.5-4.5 s−1. From determined koff rates, we conclude that once bound, the ligand will not dissociate during the transcription interval for ligand binding. SIMULATIONS OF

CO-TRANSCRIPTIONAL FOLDING We determined all relevant conformational transition kinetics along and between pathways 1, 2, and 3 by real-time NMR and stopped-flow spectroscopy, allowing us to

calculate the kinetics of the switch between different regulatory pathways (Fig. 5). We performed simulations of the conformational transition from PA–T to PAT between transcript lengths

113 and 137 at different transcription rates (0–100 nt s−1) (Supplementary Fig. 5) and derived the fraction of ON state after this transcription interval (Fig. 5a). Equilibration from the

metastable PAT-H state to the PA–TH state can be initiated once nucleotide 139 emerges from the transcribing polymerase, one nucleotide prior to the regulatory signal. Therefore, we consider

nucleotide 137 to represent the latest possible regulatory decision point. The fraction of ON state obtained from individual simulations at different transcription rates is shown in Fig. 5b

for (i) 2′dG is not bound (in green) and (ii) 2′dG is bound by 70–100% (in red) to the PA–T conformation by the time nt 113 is synthesized. Figure 5c visualizes the effective ligand-induced

kinetic delay of PT formation at a transcription rate of 20 nt s−1 that offsets equilibrium populations previously determined15. Kinetic traces between nucleotides 113 and 137 correspond to

individual simulations derived from Supplementary Fig. 6. In the presence of ligand, the simulations further distinguish between OFF state (pathways 1 + 3) and ligand-bound state (pathways

3), where the OFF state includes ligand-free folded aptamer domains that also terminate transcription. The simulations show that the riboswitch functions only at transcription rates of 10–50

 nt s−1, a value that matches those typical for bacterial polymerases (Fig. 5b). Slow transcription rates <10 nt s−1 extend the time interval for ON state folding to a point, where ligand

binding can no longer delay ON state folding sufficiently. At transcription rates exceeding 70 nt s−1, ON state folding in both absence and presence of ligand cannot compete with the rate

of transcription. In this system of competing kinetic requirements, a maximum regulation efficiency of 40–50% is achieved only at transcription rates of 10–50 nt s−1. Comparison of static

and kinetic experiments shows that the equilibration kinetics of metastable states neither lead to complete formation of the functional ON state from pathway 1 or preservation of the

functional OFF state in pathway 3 during transcription. IN VITRO TRANSCRIPTION ASSAYS We performed single-round in vitro transcription assays in the absence and presence of ligand using

both, _E_. _coli_ and _B_. _subtilis_ DNA-dependent RNA polymerases (RNAP) to (i) experimentally validate that the riboswitch functions only in a defined window of transcription rates and to

(ii) assess whether the co-transcriptional folding pathway of the riboswitch is sensitive to the transcription properties of the polymerases from different organisms. We determined the

dependence of transcription rates on temperature and NTP concentrations. We further determined the regulation efficiency by quantifying the amount of 235 nt fragments elongated past the

terminator sequence. For both polymerases, three pause sites (PS1) were determined in time-resolved transcription reactions. The first PS1 is encountered when the aptamer is fully

transcribed (Fig. 6a, Supplementary Fig. 7). A decrease in temperature from 37 °C to 4 °C increases the _t_1/2 value of PS1 from 3 s to 50 s in the absence of ligand for _E_. _coli_ RNAP.

The second PS1 is located around nucleotide 116 at the 3′-end of segment T. This second PS1 pauses the transcription complex longer than the first PS1 (24 s at 37 °C). The third PS1 is

located at the translation start site. This PS1 is located beyond the regulatory decision point and is, thus, not relevant for the co-transcriptional folding pathway of the riboswitch. For

the _E_._ coli_ RNAP, the ligand-dependent ON state fraction is primarily affected by changes in temperature, while variations in the NTP concentration in the range of 0.05–2.3 mM do not

affect the transcription rate sufficiently to cause changes in the co-transcriptional folding pathway (Supplementary Fig. 6). In the absence of ligand, the fractions of ON state represent

49–61% of all transcripts, and slightly increases with decreasing temperature (Fig. 6b). For the highest transcription rate (15.1 ± 4.5 nt s−1, 37 °C) (Supplementary Fig. 7) the fraction of

ligand-independent ON state is 49 ± 12% and for the lowest transcription rate (1.2 ± 0.2 nt s−1, 4 °C) the fraction of ON state is 61 ± 9%. In line with riboswitch function, addition of

ligand promotes accumulation of nascent transcripts within pathway 3 and decreases the fractions of ON state obtained to 35–43% for transcription reactions performed at 37 °C and 25 °C. For

multi-round transcriptions, the effect of ligand addition on the fraction of ON state obtained is significantly more pronounced with 40–50% change in population ratio opposed to ~20% for

single-round transcriptions (Fig. 5c). For transcription reactions performed at 4 °C, ligand addition does not cause significant changes in the population of ON state for both single-round

and multi-round transcription. For the _B_. _subtilis_ RNAP, transcription rates could only be determined at 37 °C due to insufficient transcription efficiency at lower temperatures.

Transcription rates for _B_. _subtilis_ RNAP are higher compared to _E_. _coli_ RNAP (34.7 ± 30.7 nt s−1) (Supplementary Fig. 7). In addition, the decrease in ON state population upon

addition of ligand is significantly less pronounced. At an NTP concentration of 2.3 mM, ligand addition only induces a 5% shift in the fraction of ON state. Contrary to the behavior of _E_._

 coli_ polymerase, a decrease in the NTP concentration to 0.05 mM increases the ligand-induced shift in the fraction of ON state to 21%. For _E_. _coli_ polymerase, the overall regulation

efficiency determined by transcription multi-round assays matches remarkably well with values predicted by our simulations of 40–50%, provided transcription reactions were performed close to

the physiological temperature of _E_._ coli_. While an NTP-concentration range of 0.05–2.3 mM does not appear to affect transcription rates significantly to influence the co-transcriptional

folding pathway of the riboswitch, a reduction in temperature to 4 °C leads to near-complete loss of riboswitch function. Determined transcription rates clearly show a negligibly small

decrease in transcription speed from 37 °C to 25 °C, while transcription rates drop below the functional window of the riboswitch (<10 nt s−1) as determined by our simulation at 4 °C. In

agreement with our simulations, slower transcription rates at 4 °C expand the time span available for PA–T to PAT(M) folding and lead to an increase in structures populating the ON state at

the regulatory decision point in both absence and presence of ligand. In stark contrast, the _B_. _subtilis_ polymerase requires a reduction in NTP concentration to induce transcription

termination upon ligand addition. Due to the high percentage of full-length transcripts this effect is likely unrelated to ligand binding or ON state folding, but is linked to the inability

of terminator formation due to the rapid elongation by the polymerase. The calculated transcription rates from the in vitro assays are not absolute values for single RNAPs, but instead are

composite rates, as they indicate the point in time at which half of the RNAPs completed transcription of an RNA fragment of a specific length. This approximate rate can then be used in the

simulations to examine the relationships of the different parameters in the regulation mechanism. DISCUSSION To act as transcriptional regulatory elements, the sequence of riboswitches has

to be optimized to fulfill a number of kinetics requirements. Our experimental approach delineates the competition between refolding rates of cis-regulatory RNA and processive transcript

elongation. Within the regulatory decisive 2′dG concentration range of 0.1–100 µM for dGsw26, ligand association rates for purine aptamer domains of ~104–105 M−1 s−1 27,28 can effectively

direct ligand-binding to either compete with or lag behind bacterial polymerases synthesizing the relevant ligand-binding sequence at rates of 0.5–4.5 s−1. Remarkably, PS1 induces

transcriptional stalling at the latest possible transcript length with highest ligand binding affinity according to previous results29. To determine shifts in population ratios as

transcription proceeds further, we present a ligation-based approach that provides access to reversible trapping of metastable states that form within the activation (pathway 1) and

repression pathway (pathway 3) of ON function (pathway 2). In the biological context of co-transcriptional folding, single nucleotides are sequentially released to an ensemble of RNA

intermediates differing in length that are transcribed at locally varying transcription rates. Real-time monitoring of such folding is experimentally challenging and cannot be accomplished

by established methods. In the approach applied here, folding is monitored by immediately releasing a part of the riboswitch expression platform to interact with the aptamer domain. Despite

these experimental limitations, we have selected a representative transcription intermediate that most accurately reflects refolding rates of the metastable state on average to obtain

high-quality quantitative data. We linked kinetics derived from light-induced release of the stable ON state to unbiased mapping of conformational space15 to assess the kinetics of

ligand-dependent conformational switch. We further experimentally validated the impact of transcription rates and polymerase dependent transcription properties on the regulation efficiency

of the riboswitch using _E_._ coli_ and _B_. _subtilis_ RNAP. Our results demonstrate the requirement for a strict synchronization of transcription intervals of metastable states,

corresponding transcription rates, and refolding rates, where transcription elongation must not only be matched to refolding and ligand binding kinetics but also to folding delays induced by

thermodynamic stabilizations to preserve metastable states within distinct sequence intervals. A significant offset in temperature from physiological conditions leads to quantitative

folding to the ON state from both metastable ligand-bound and ligand-free aptamer domains. Sequence-specific variations in transcription rates impact both ON state folding from pathways 1

and 3 equally. Therefore, it is critical that pausing within the expression platform is adapted to the sequence-specific transcription speed of the native polymerase. In a previous study we

could already show that the detailed investigation of transcriptional riboswitches requires the native RNA polymerase30. Here, we further substantiate this proposal by revealing the

difference in co-transcriptional riboswitch folding transcribed by two non-native polymerases. Furthermore, biologically demanding experiments with _M_. _florum_ polymerase would be required

to compare these results to native 2′dG riboswitch folding. General fluctuations in transcription rates resulting from changes in cellular conditions31 may be self-regulated by the negative

feedback loop of the riboswitch itself. The kinetic and simulation framework developed here provides an important contribution to the conceptual basis to understand transcriptional

riboswitches. METHODS SAMPLE PREPARATION dGsw85 and mutated dGswC74U constructs with varying length for stopped-flow spectroscopy were prepared by in vitro transcription from PCR products by

adjusting the reverse primers accordingly as reported previously32. For dGswC74U constructs, two artificial G’s were incorporated at the 5′-end in the forward primers to enable rapid RNA

purification by buffer exchange. dGsw85 was transcribed with a 5′-hammerhead ribozyme, exchanged into water, and directly used for ligation reactions. Photocaged dGsw86–121 was prepared via

solid phase synthesis in a 17 µmol scale. Ultramild phosphoramidites, chemical phosphorylation reagent I, and (_S_)-NPE-U phosphoramidite (Supplementary Fig. 2) were used and a postsynthetic

DEA wash performed. The use of NH3:EtOH (3:1) for deprotection primarily generates a byproduct with a mass 53 Da greater than the desired product (Supplementary Table 1). The dGsw86–121

fragment could be isolated using 3 mL _t_-BuNH2:water (1:3) for 6 hours at 60 °C. The mixture was separated in 10 reaction tubes and the solvent evaporated. Each tube was treated as follows:

the 2′-deprotection was performed using 6.5 mL of a mixture of NMP:Et3N:Et3N·3 HF (3:1.5:2) for 3 hours at 60 °C. After addition of 30 mL _n_-BuOH the mixture was stored at −80 °C for 2 

hours and then centrifuged at −8 °C for 1 hour. The supernatant was decanted and the residue purified via anion exchange HPLC (Amersham Bioscience with an UV-900 detector and P-900 pumps)

using a Tricorn SOURCE 15Q AEX 10/150 column (gradient: 0–100% 0.35 M LiClO4 in water in 17 column volumes, flow: 4 mL min−1). The collected oligoribonucleotide was desalted using a Tricorn

SOURCE 15 RPC 10/150 column (gradient: 5–40% MeCN in 0.1 M TEAA buffer pH 7 in 9 column volumes, flow: 4 mL min−1). The identity of the oligoribonucleotide was confirmed by ESI-MS: mass

calculated: 11784.5 Da, found: 11785.7 Da. Photocaged dGsw121 was prepared by enzymatic splinted ligation using T4 RNA ligase 2. Fragments dGsw85 (15N-G/U) and photocaged dGsw86–121 were

preannealed to a 31 nt splint (splint:31AATAATTAAAGAGTGGTTTTTCACTTATAGT) prior to ligation at 25 °C for 15 min. Ligation reactions were performed with 12.5 µM dGsw85, 6.25 µM caged

dGsw86–121, 10 µM splint31, 48 µg mL−1 T4 RNA ligase 2 (homemade), 10 mM MgCl2, 2 mM ATP, 25 mM DTT in 50 mM Tris-Cl, pH 7.4. dGsw85 was applied in 2-fold excess to optimize the turnover for

photocaged dGsw86–121 (80%). Ligation reactions were incubated at 37 °C for 2 hours followed by overnight incubation at 25 °C, followed by incubation with TURBO DNAse (0.02 U µL−1) (Thermo

Fisher Scientific) for 8 hours. Ligation reactions were directly applied to HPLC using a Kromasil RP18 10 × 250 column without further concentration and eluted at room temperature (gradient:

0–40% MeCN in 0.1 M TEAA, pH 6.0). Purified photocaged dGsw121 was lyophilized and exchanged into NMR buffer (50 mM KCl, 25 mM K2HPO4/KH2PO4, pH 6.2). Final yield of photocaged dGsw86–121

was 0.6 µmol (54 %). NMR-SPECTROSCOPY Time-resolved NMR experiments were performed as reported previously17,18,33 in a pseudo-2D experiment using the jump return echo water suppression

scheme34. Laser irradiation within the NMR tube was triggered via a TLL connection to a laser set up (Paladin Advanced 355-8000). Kinetics of ligand-free samples were recorded after 500 ms

and kinetics of ligand-bound RNAs after 1 s of laser irradiation. The signal intensity was maximized by Ernst angle excitation35. Kinetics traces of ligand-free samples were averaged over 10

individual experiments and of ligand-bound samples over two individual experiments on 100 µM caged dGsw121 samples. 1H,15N-BEST-TROSY experiments were recorded with modifications according

to Brutscher et al36,37. NMR experiments were performed on a Bruker 800 MHz spectrometer equipped with a 5-mm _z_-axis TCI-HCN cryogenic probe. For processing of data, Topspin 3.5 (Bruker

Biospin) was used. STOPPED-FLOW SPECTROSCOPY Stopped-flow measurements were performed on a π*-180 CDF-spectrometer (Applied Photophysics). Folding was monitored by rapid mixing of dGswC74U

(2 µM) with 2-aminopurine 2′-deoxyriboside (2 µM–512 µM) in a 1:1 volume ratio at 25 °C. Both ligand and RNA were provided in NMR buffer with a Mg2+ concentration of 3 mM. 2′dAP was excited

at 304.5 nm using 2 mm slit widths and fluorescence was monitored at >340 nm. Kinetic traces were averaged over 10–15 single measurements. Fluorescence decays were fitted with single and

bi-exponential decays with three and five parameters, respectively (Supplementary Fig. 4). Association (kon) and dissociation (koff) constants were determined from individual apparent rate

constants according to equation (1) (Supplementary Table 3): $$k_{{\rm {app}}} = {k_{\rm {on}}} \times \left[ {2{\prime}\mathrm{dAP}} \right] + {k_{{\rm {off}}}}$$ (1) SIMULATIONS OF

CO-TRANSCRIPTIONAL FOLDING Simulations of co-transcriptional folding were performed by kinetic Markov simulations. Simulations were performed as described previously15, starting from the

aptamer domain fold at transcript length of 113 nt, followed by simultaneous elongation to a transcript length of 137 nt at transcription rates of 10–100 nt s−1 and folding to the

antiterminator conformations PAT(M) and PAT(I) with rate constants determined by real-time NMR. TIME-RESOLVED TRANSCRIPTIONS The DNA template used for _E_. _coli_ and _B_. _subtilis_ RNAP

mediated transcription was composed of the lambdaPR promoter followed by the dGsw-coding sequence

(GCAAATCGCGCTGTTAGCGGGCccttgactattttacctctggcggtgataatggttgcATGAATATAAAAGAAACTTATACAGGGTAGCATAATGGGCTACTGACCCCGCCTTCAAACCTATTTGGAGACTATAAGTGAAAAACCACTCTTTAATTATTAAAGTTTCTTTTTATGTCCAAAAGACAAGAAGAAACTTTTTTATTTAGTTGAATTTATAATAAGAGAAAAAGAAAGGATATTATATGGCAAAAATAAAAAACCAATATTACAACGAGTCTGTTTCGCCAATTG).

The time-resolved transcriptions were performed as described previously30 at 37 °C, 25 °C, and 4 °C with NTP concentrations of 0.05 mM, 0.1 mM, 1 mM, and 2.3 mM. Transcription rates were

determined as described previously38 and PS1 were analyzed as described by Landick 199639. DATA AVAILABILITY The data that support the findings within study are available from the

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leader pause site as paradigm. _Methods Enzymol._ 274, 334–353 (1996). Article  CAS  PubMed  Google Scholar  Download references ACKNOWLEDGEMENTS This work was supported by DFG (CRC 902 and

GRK 1986 “CLiC”), and Fonds der Chemischen Industrie (C.H.). H.S. and A.H. are members of the DFG-funded cluster of excellence: dynamics of macromolecular complexes (EXC 115). BMRZ is

supported by the state of Hesse. AUTHOR INFORMATION Author notes * These authors contributed equally: Christina Helmling, Dean-Paulos Klötzner, Alexander Heckel. AUTHORS AND AFFILIATIONS *

Institute for Organic Chemistry and Chemical Biology Center for Biomolecular Magnetic Resonance (BMRZ), Johann Wolfgang Goethe-Universität, Max-von-Laue-Straße 9, 60438, Frankfurt, Germany

Christina Helmling, Florian Sochor, Anna Wacker, Boris Fürtig & Harald Schwalbe * Institute for Organic Chemistry and Chemical Biology, Johann Wolfgang Goethe-University Frankfurt,

Max-von-Laue-Straße 9, 60438, Frankfurt, Germany Dean-Paulos Klötzner & Alexander Heckel * Department of Biochemistry, University of Wisconsin–Madison, Madison, WI, 53706, USA Rachel

Anne Mooney & Robert Landick Authors * Christina Helmling View author publications You can also search for this author inPubMed Google Scholar * Dean-Paulos Klötzner View author

publications You can also search for this author inPubMed Google Scholar * Florian Sochor View author publications You can also search for this author inPubMed Google Scholar * Rachel Anne

Mooney View author publications You can also search for this author inPubMed Google Scholar * Anna Wacker View author publications You can also search for this author inPubMed Google Scholar

* Robert Landick View author publications You can also search for this author inPubMed Google Scholar * Boris Fürtig View author publications You can also search for this author inPubMed 

Google Scholar * Alexander Heckel View author publications You can also search for this author inPubMed Google Scholar * Harald Schwalbe View author publications You can also search for this

author inPubMed Google Scholar CONTRIBUTIONS C.H., D.-P.K., F.S., and R.A.M. performed experiments. D.-P.K. conducted the chemical synthesis of the photocaged RNA. F.S., and R.A.M.

performed transcription assays. All other experiments were conducted by C.H. The contributions by C.H. and D.-P.K. were both essential for the work, C.H. and D.-P.K. should be regarded as

joint first authors. All authors contributed to the design of the experiments, analysis of data, discussion, and writing of the manuscript. CORRESPONDING AUTHORS Correspondence to Alexander

Heckel or Harald Schwalbe. ETHICS DECLARATIONS COMPETING INTERESTS The authors declare no competing interests. ADDITIONAL INFORMATION PUBLISHER'S NOTE: Springer Nature remains neutral

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al._ Life times of metastable states guide regulatory signaling in transcriptional riboswitches. _Nat Commun_ 9, 944 (2018). https://doi.org/10.1038/s41467-018-03375-w Download citation *

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