High-throughput, fluorescent-aptamer-based measurements of steady-state transcription rates for the Mycobacterium tuberculosis RNA polymerase

Abstract The first step in gene expression is the transcription of DNA sequences into RNA. Regulation at the level of transcription leads to changes in steady-state concentrations of RNA transcripts, affecting the flux of downstream functions and ultimately cellular phenotypes. Changes in transcript levels are routinely followed in cellular contexts via genome-wide sequencing techniques. However, in vitro mechanistic studies of transcription have lagged with respect to throughput. Here, we describe the use of a real-time, fluorescent-aptamer-based method to quantitate steady-state transcription rates of the Mycobacterium tuberculosis RNA polymerase. We present clear controls to show that the assay specifically reports on promoter-dependent, full-length RNA transcription rates that are in good agreement with the kinetics determined by gel-resolved, α-32P NTP incorporation experiments. We illustrate how the time-dependent changes in fluorescence can be used to measure regulatory effects of nucleotide concentrations and identity, RNAP and DNA concentrations, transcription factors, and antibiotics. Our data showcase the ability to easily perform hundreds of parallel steady-state measurements across varying conditions with high precision and reproducibility to facilitate the study of the molecular mechanisms of bacterial transcription.


INTRODUCTION
Cellular RNA abundance is dictated by the relati v e steadysta te ra tes of RNA production and degradation.In particular, the rate of RNA production is ubiquitously the target of gene regulatory mechanisms and often represents a good pr oxy for pr otein synthesis flux ( 1 ).More specifically, the ra te a t w hich full-length RN A tr anscripts are gener ated is typically controlled by the rate of transcription initiation.This is because the overall initiation rate is often slower than subsequent elongation and termination steps, and because m ultiple RN A pol ymerases (RN APs) may be elongating at the same locus at the same time.Bacterial transcription initiation pr ogresses thr ough se v eral intermediates, where the rates and equilibrium constants that describe the initial binding of the RNAP to the promoter and the subsequent isomeriza tion steps tha t culmina te in opening of promoter DNA can vary greatly depending on the sequence of the e99 Nucleic Acids Research, 2023, Vol. 51, No. 19 PAGE 2 OF 16 promoter ( 2 , 3 ).For example, Esc heric hia coli promoters can differ in rates of promoter opening by factors of 10 3 -10 4 , resulting in initiation e v ents ranging from once per second to once per generation ( 4 , 5 ).Following promoter opening, binding of the first two initiating nucleoside triphosphates (NTP) forms an initially transcribing complex that begins producing a nascent RNA transcript.After formation of the initiating dinucleotide, each step of RNA synthesis requir es translocation, str essing RNAP-promoter contacts ( 6 , 7 ).If this stress disrupts these contacts, the RNAP escapes from the promoter and begins processi v e elongation.Otherwise, complexes stall near the promoter or perform cy cles of aborti v e transcription ( 5 , 8 , 9 ).After decades of careful biophysical dissection, dri v en mainly by pre-steadystate kinetic and structural biology approaches, bacterial initia tion pa thways are well-characterized on a handful of different promoters for model bacteria (re vie wed in ( 2 , 3 , 10-12 )).Many kinetic / structural intermediates have been identified, including of f-pa thway sta tes tha t, in some cases, do not lead to full-length RNA production ( 8 , 13-17 ).The resulting models from all these studies are complex and can vary depending on the system studied (18)(19)(20), bringing into question model generalizability across bacteria and different promoter sequences.
Ideally, rate constants for RNAP binding, DNA opening, initial transcription and promoter escape obtained from pre-stead y-sta te kinetic measurements could be used to calculate an overall initiation rate or the average time between initiation e v ents.A demonstration of this can be seen in a pproaches w here both basal and regulated RNA flux is calculated based on simple models of transcription initiation ( 3 , 21 ).Howe v er, while these models may address theoretical links between initiation kinetics and steady-state rates of RNA production, they cannot account for the large number of variables that determine the rate of transcription .This overall initia tion ra te is most simply described using Michaelis-Menten enzyme kinetics, where the RNA polymerase is the enzyme, promoter DNA is the substrate, and the full-length RNA transcript is the product ( 22 ).This model assumes a constant concentration of RNAPpromoter (enzyme-substrate) complexes generated by a balance between promoter binding and escape rates and predicts a constant reaction velocity or rate of transcript production.Within this formalism, regulated promoters are activated or repressed via changes in the Michaelis-Menten parameters K m (the concentration at which the half maximal rate is observed) and / or V max (the maximal rate observed under saturating conditions) without changes in the free RNAP concentra tion.Alterna ti v ely, constituti v ely acti v e promoters are affected by cell growth-ratedependent changes in the free RNAP concentration, independent of changes in K m and / or V max (23)(24)(25).Classic examples of environmental adaptation in bacteria affecting transcription initiation processes include the upregulation of beta-galactosidase in response to the presence of lactose and the absence of glucose ( 26 ), and the genomewide response to starvation known as the stringent response ( 27 , 28 ).
In vitro measurements of steady-state transcription rates can be used to test and de v elop more comple x models but have been limited by current methodolo gies.Historicall y, measurements of in vitro basal and regulated transcription initiation kinetics have been made possible by monitoring RNA production via the incorporation of radio-labelled NTPs, resolved via polyacrylamide gel electrophoresis ( 29 ).The approach was used in the initial discovery of RNAP holoenzyme activity more than fifty years ago ( 30 ) and is the most common assay for probing transcriptional mechanisms in vitro .The strengths of the assay include its high sensitivity and single-nucleotide resolution e v en with small quantities of RNA.This allows for the separation of RNA products by length and to inv estigate indi vidual steps in nucleotide addition reactions that underlie the fundamental mechanisms of RNAP.Howe v er, stead y-sta te 32 P-detected transcription assays have several drawbacks.A practical limita tion is tha t the use of radioactivity is e xpensi v e, both for the purchase of the reagent itself and for the requirement of specialized protocols for safe shipping and use in the lab.The reaction must also be taken through se v eral steps that may include the incorporation of proteases or chelators to quench the reaction at set timepoints, phenol / chloroform extr actions, and / or denatur ation and loading the sample on an electrophoretic polyacrylamide gel.All these steps, as well as the quantification of resolved product bands via image analysis, can introduce non-biochemical variability in the measured RNA amounts.In addition, these assays r equir e significant amounts of time and training, where single experiments often take multiple da ys bef ore complete quantification and where technical practice is needed for generating reproducible data.All these factors combined result in relati v ely limited throughput.As a result, the timedependent measurements needed to quantitate steady-state initia tion ra tes ar e infr equently performed and single timepoint measurements are often used to infer mechanisms of gene regulation.
In contrast to the in vitro approaches described above, in vivo transcription studies have undergone a remarkable transformation where genome-wide transcript le v els in cells under varying environmental and genetic backgrounds are routinely queried ( 31 ).We were motiv ated b y the desire to increase the throughput of in vitro transcriptional studies to facilitate the efficient testing of mechanistic hypotheses, the de v elopment of predicti v e transcription initiation models, and to be able to compare transcript flux from reconstituted systems to the genome-wide information accessible via RNAseq.To this end, we explored the use of a fluorescent-aptamer-based detection system to report on in vitr o stead y-sta te tr anscription r ates.
Fluorescent light-up aptamers are RNA sequences that bind small molecule fluorophore dyes and generate a large fluorescence enhancement.They have been used in numerous applications (re vie wed in ( 32 )) including the detection of nascent RNA transcripts in cells (33)(34)(35)(36)(37)(38), in synthetic biology tr anscription-tr ansla tion coupled in vitr o systems for T7 RNAP (39)(40)(41) and with cell lysates from di v erse bacterial species ( 42 ).These a pproaches rel y on the use of a DNA template that encodes for an aptamer sequence so that each transcription e v ent results in the generation of an RN A a ptamer w hich folds and binds the dye.For each transcript produced, there is an accompanying incr ease in fluor escence (Figur e 1 ).Importantly, as the change in dye fluorescence requires the synthesis of a full-length RNA transcript containing the aptamer, the fluorescent readout is not complicated from short aborti v e products that may be generated during promoter escape.Using this approach, recent work has illustrated how the rates of in vitro reactions can be quantified in a fluorimeter cuvette with E. coli RNAP ( 43 ) and in a plate-reader format with T7 RNAP ( 44 ).Here, we follow up on these studies and provide a description of essential control experiments needed to clearly link the fluorescent signal with the transcription of a promoter-deri v ed product.Additionally, using data acquired with RNAP from Mycobacterium tuberculosis ( Mtb ), we describe a workflow for the calibration and quantification of RNA concentration in multi-round initiation kinetics analyzed with a Michaelis-Menten approach.Most significantly, we illustrate the utility of this approach for the investigation of mechanisms of transcriptional regulation by NTP concentr ation, tr anscription factors, and antibiotic inhibitors, all deri v ed fr om high-thr oughput measurements under stead y-sta te conditions.

Reagents
Pr epar ation of DNA constructs.Circular plasmid templates, 2557 base-pair (bp) in length, were ordered from Twist Bioscience (San F rancisco , CA), containing the sequences corresponding to the Mtb ribosomal RNA promoter ( rrnA P3), the Spinach-mini aptamer ( 33 ) and the E. coli rrnB P1 T 1 terminator ( 45 ) (illustrated in Figure 1 ).Plasmids were isolated via the Qiagen Midi Prep Kit (catalogue #: 27106) and were predominantly supercoiled (Supplementary Figure S1).For additional sequences and further descriptions, see Supplementary Table S1.Linear PCR templates 250 bp in length were amplified from 200 nucleotide (nt) PAGE purified DNA Ultramer oligos (Integrated DNA Technologies, Inc., Coralville, IA) following annealing and end-filling.For more details including primer and final construct sequences, as well as further preparation details, see Supplementary Table S2.

Plate-r eader fluor escence measur ements
To measure multi-round kinetics under stead y-sta te conditions in real-time, we monitored the change in DFHBI fluorescence upon binding to a transcribed, full-length RNA sequence containing the Spinach-mini aptamer.Data was collected using a Synergy2 multi-detection microplate reader (BioTek Instruments, Inc., Winooski, VT) with the corresponding Gen5 analysis software.DFHBI fluorescence was measured with a tungsten light source equipped with a 505 nm long-pass dichroic mirror, excited with a 460 ± 40 nm bandpass filter, and the resulting emission signal was monitored with a 528 ± 20 nm bandpass filter.Data was acquired at a read height of 7.00 mm, typically in 20-30 s intervals, with varying total acquisitions times, not exceeding 75 min.
Transcription reaction master mixes containing 90% of the final volume included the RNAP holoenzyme , NTPs , DFHBI, and RNase inhibitors.Based on the volumes added for each corresponding buffer addition and concentrated stock component, the final solution conditions were 20 mM Tris (pH 8.0 at 25 • C), 40 mM NaCl, 75 mM Kglutamate, 10 mM MgCl 2 , 5 M ZnCl 2 , 20 M EDTA, 5% (v / v) glycerol (defined as transcription buffer) with 1 mM DTT and 0.1 mg / mL BSA.From these master mixes, a volume of 9 l for each technical replicate was transferred to an individual well in a 384 well, lo w v olume, round-bottom, non-binding polystyrene assay plate (Corning; catalogue # 4514).For the negati v e controls, which r epr esent the entirety of the reaction components except for DNA, 1 l of transcription buffer was added to account for the remaining 10% of the final reaction volume (10 l).Wells were covered with an optical adhesi v e to pre v ent sample evaporation (A pplied Biosystems; catalo gue # 4360954).The plate was then incubated for 10 min at 25 • C, followed by a 30 s shake agitation in the microplate reader.An initial reading of the negati v e controls was used to scale the automatic gain adjustment to the background signal with an arbitrary value of 1000 RFUs and was applied to all the subsequent reads.The adhesi v e cov er was remov ed, and the transcription reaction was initiated with 1 l DNA unless otherwise indicated.Multichannel pipettors were used to reduce the initiation time difference across wells.Once the DNA was added, the plate was agitated for 15 sec before starting the kinetic measurements.Unless otherwise stated, the reaction master mixes always contained 20 M DFHBI and 0.4 U / l RNase inhibitors.For additional details regarding reaction specifics for individual titrations, including titrations measured under single-round conditions, see Supplementary Materials and Methods.

Radio-labelled NTP incorpor ation tr anscription experiments
Transcription experiments were performed with 500 M each NTP under identical solution conditions and temperature as multi-round fluorescence experiments.Mtb A RNAP holoenzyme and DFHBI concentrations were varied and always included unless otherwise indicated.20 nM of ␣-32 P UTP was added to label the nascent RNA transcripts via incorporation by RNAP.Reactions were initiated by addition of circular plasmid constructs containing either 5 nM Mtb rrnA P3 or a template lacking a promoter (promoterLESS, see Supplementary Table S1).5 l aliquots wer e r emoved from the r eaction mixtur e at 1, 5, 10, 15, 20 and 30 min and combined with 5 l of quench solution containing 95% formamide, 0.015 M EDTA, 0.05% (w / v) xylene cyanol, and 0.05% (w / v) bromophenol blue.5 l of each quenched timepoint was then loaded onto a 5% denaturing PAGE gel.Gels were run in TBE for 2 to 3 h at 1,500 V and then transferred to a phosporimaging cassette.After exposing for 18 h, phosphorimaging scr eens wer e imaged via a Typhoon 9000 phosphorimager.Bands of interest were quantified using the ImageQuant software and converted to reaction RNA concentrations as previously described ( 5 ).

Analyses of fluorescent time courses
Extraction of steady-state rates.As the fluorescent signal in the absence of aptamer formation does not significantly change across conditions w here DN A or NTPs were left out of the reaction (Supplementary Figure S3), the same negati v e control could be used to correct all experimental conditions within a titration.This applies only if the time course measurements are made under the same solution conditions and instrumental detection scaling.Howe v er, to minimize e xperimental and / or instrumental variation, each time a ne w e xperiment was performed, a minimum of 2 to 3 technical replicates of the negati v e control (leaving out DNA) were collected and measured concurrently with the experimental data.Prior to data analysis, the experimental traces underwent two subtractions: first, the fluorescent value recorded at the initial timepoint was subtracted from all timepoints, bringing the starting fluorescent value to zero, and second, the fluorescence from the experimental traces was subtracted using the corresponding signal from each time-point of the negati v e control.
Linear r egr ession of the corr ected fluor escent traces was performed with a custom MATLAB fitting program (described further in Results).Using a statistical weighting from multiple technical replicates per condition and a user inputted R 2 value to define the goodness of fit (Supplementary Figure S6), an unbiased determination of the linear regime can be obtained, where the slope of the fitted line reports on the stead y-sta te ra te in units of change in fluorescence / time.In the work pr esented her e, R 2thresholds > 0.998 were used.

Non-linear r egr ession analyses of concentration dependencies.
For analyses of titration data that yielded hyperbolic concentration dependencies in steady-state rates, a Michaelis-Menten equation was applied, fitting the data to Equation 1 where v represents the stead y-sta te ra te and S r epr esents the titr ated substr ate, either DNA or individual NTPs.Steadysta te ra tes determined in some NTP titrations were not welldescribed by Equation 1, displaying sigmoidal concentration dependences.These titrations were fit to Equation 2, a modified Michaelis-Menten equation with three parameters: V max , an apparent K m ( K m , app ), and the exponent n .
Here, we define K m , app as the NTP concentration that results in half maximal velocity, which is equivalent to the PAGE 5 OF 16 Nucleic Acids Research, 2023, Vol. 51, No. 19 e99 standard operational definition of K m when n = 1.It should be noted that while this equation is functionally identical to the Hill equation, if a non-unity n is the result of the kinetics from a single-site-binding system, then n is only effecti v ely demonstrati v e of the type (positi v e / negati v e) and magnitude of cooperativity.In such a case, n has no specific physical meaning as it does in more traditional cases such as cooperati v e binding of multiple ligands ( 49 ).The kinetics of transcription inhibition as a function of antibiotic concentra tion a t constant NTP concentra tions were also found to be sigmoidal.For determination of the half maximal inhibitory concentration ( IC 50 ) of Rifampicin and Fidaxomicin, the concentration-dependent reduction in steadysta te ra tes was fit to Equa tion 3 .

Statistical analyses
For all fluorescent data presented unless otherwise indicated, between 2 and 4 independent experiments were collected for each condition tested.Within each independent e xperiment, standar d de viations from 2 to 3 technical replicates were used as a statistical weight during the linear regression analyses.All non-linear regression analyses were performed using the standar d de viations of the independent experiments as a statistical weight.Errors in V max , K m,app , n and IC 50 values are those obtained from the fit of the averaged data set.For simplicity, only a single r epr esentati v e independent e xperiment is shown in the plots.For gelbased quantifications, average values and standard deviations from 2 to 3 independent experiments are reported.

Data availability
The MATLAB code used for automated linear r egr ession analyses has been made publicly available and can be accessed with the following GitHub link ( https://github.com/egalburt/aptamer-flux-fitting ).

Promoter -dependent, aptamer -based measur ements of fulllength transcription
As similar assays have been described elsewhere, we briefly describe the specifics of the version we utilized for these studies.DNA templates were constructed such that an 80 nt Spinach-mini aptamer sequence ( 33 ) was inserted 30 bp downstream of the +1 transcription start site corresponding to the genomic Mtb ribosomal RNA promoter sequence ( rrnA P3).Specifically, -60 to +31 of rrnA P3 was included to account f or an y upstream-promoter interactions as well as the initially transcribed sequences, both of which can regula te initia tion kinetics.Inclusion of the genomic initially tr anscribed sequence, r ather than just positioning the aptamer sequence to start at the +1 position, provides for unaltered promoter esca pe, w hich is known to become rate limiting under certain conditions (see ( 3 ) for examples of sequence effects on initiation kinetics).The E. coli intrinsic rrnB P1 T 1 terminator sequence was included in circu-lar plasmid DNA templates to dissociate the ternary polymerase , template , and transcript complex.Sequences for both circular plasmid and linear PCR templates are gi v en in Supplementary Tables S1 and S2.
When transcribed downstream of the Mtb rrnA P3 promoter, the Spinach-mini aptamer sequence folds and binds the small molecule fluorophore DFHBI, resulting in a fluorescence enhancement.By mixing RNAP, DNA, all NTPs and DFHBI, the reaction is monitored in real-time wher e the incr ease in fluor escence r eports on transcript production.All experiments were performed in a 384 well pla te-reader forma t using 10 l reaction volumes facilitating high-throughput measurements of stead y-sta te initiation kinetics with minimal sample volume r equir ements.The slope of the fluorescent signal at early times reports on the stead y-sta te ra te of transcription initia tion (Figure 1 ).In general, one expects initiation kinetics and not cotranscriptional aptamer folding ( 39 , 50 , 51 ) or dye binding ( 52 ) to be rate-limiting for functional aptamer production.Howe v er, e v en when the initia tion ra te is not limiting relati v e to the timescales of elongation or aptamer folding, the stead y-sta te ra te will still specifically report on the initiation rate (Supplementary Figure S2).This is because once one RNAP leaves the promoter, another RNAP may bind and begin the process of initiation irrespecti v e of these downstream processes.This condition may be broken by patholo gical cases w her e downstr eam pausing cr ea tes a traf fic jam that backs up onto the promoter, influencing the time needed for the next polymerase to bind the promoter.
Transcription reactions were initiated by adding DNA templates to Mtb RNAP preincubated with NTPs, DFHBI dye and RNase inhibitors.Fluorescence was monitored in r eal-time, wher e we observed three distinct kinetic phases to the trace using the circular plasmid template containing the rrnA P3 sequence: (i) a lag time, (ii) a linear increase and (iii) r oll-off fr om the linear regime that begins to plateau over time (Figure 2 A, black curve).We note that when starting fr om a well-contr olled time zer o, the lag time theoretically reports on the time required for pioneering RNAPs to complete the initiation process, transcribe the aptamer, and for dye to bind the folded RNA transcript ( 43 ) (Supplementary Figure S2).In our assay setup, since initiation of the reaction was done by hand, we do not attempt to quantitate the la g time, b ut focus on the linear regime.In the absence of either DNA or NTPs, only a slow decay in DFHBI fluorescence was observed as a function of time (Supplementary Figure S3), indicating that the increase in fluorescence when all reaction components are present is due to transcriptionderi v ed aptamer formation.For all plots shown, the signal in the absence of transcription is subtracted from the experimental traces so that the y-axis reports specifically on the fluorescence generated by transcription of the aptamer (Materials and Methods).Correcting for this timedependent fluorescent change of the background signal is especially important when evaluating conditions of minimal tr anscript gener ation (Supplementary Figure S3).
To confirm that transcription is dri v en by the rrnA P3 promoter sequence, we performed the assay using a circular plasmid template lacking the pr omoter (i.e.pr omoterLESS; for sequence, see Supplementary Table S1) as a negati v e control.The signal generated in the presence of rrnA P3 is more than 20-fold higher than that obtained with the pro-moterLESS template (Figure 2 A, red), confirming that the signal arises from promoter-dri v en transcription.Howe v er, the amount of signal generated by the promoterLESS template is not zero, as can be seen by comparing to a trace using a templa te tha t contains the promoter but lacks the aptamer sequence (Figure 2 A, blue).This comparison suggests that e v en in the absence of a promoter sequence, some basal le v el of aptamer is transcribed (see below for further discussion).

Fluorescence can be monitored under both multi-and singleround conditions
Our specific goal was to obtain measurements of steadysta te ra tes of transcription.We ther efor e needed to confirm that the observed signal is due to multiple rounds of transcription.We expect significantly higher signal amplitude under multi-round conditions compared to single-round conditions, w here DN A tra ps are used to pre v ent RNAP r ebinding.The pr eincubation of the r eaction with salmonsperm DN A, w hich does not acti v el y dissociate RN APs from the DNA like other competitors such as heparin ( 20 ), resulted in the absence of any signal upon the addition of NTPs (Supplementary Materials and Methods and Supplementary Figure S4A).This result shows that the competitor DNA acts as an effecti v e trap for the Mtb RNAP and that its inclusion establishes single-round conditions.Upon initiating the transcription of pre-bound RNAP-DNA complexes with 500 M NTPs and salmon-sperm DNA competitor, we observed no lag time, and the fluorescence ra pidl y reached sa tura tion as RNAP re-binding was pre v ented (Figur e 2 B).These r esults ar e consistent with pr evious r eports of single-round conditions for full-length transcript production ( 9 , 14 , 16 ).In the absence of competitor under otherwise identical conditions, we observed a much larger increase in the overall fluorescent signal (Figure 2 B), confirming that in the absence of DN A tra p, the assay is multi-round.
In theory, use of this assay under single-round conditions with pre-formed RN AP-DN A complexes should permit kinetic analyses of processes that are difficult to de-termine in multi-round conditions, such as promoter escape.Titrating all four NTPs in the presence of the DNA trap resulted in an NTP-concentration-dependent increase in signal amplitude, where the traces could be well-fit by a single-exponential function (Supplementary Figure S4B).This NTP-concentration-dependent change in amplitude suggests that the rate of RNAP dissociation from the promoter is on the same order as the rate of promoter escape under these conditions.This observation is consistent with our previous transient-state kinetic measurements of Mtb promoter escape kinetics, where we observed that increasing NTP concentration stabilizes the RN AP-DN A complexes, slowing the rate of dissociation and facilitating escape ( 20 ).These results illustra te tha t this fluorescent-aptamer-based assay can be used under single-round conditions to examine the kinetics of sub-steps in initia tion (i.e.dissocia tion and promoter escape).

Extracting steady-state rates fr om real-time, multi-r ound fluor escent tr aces
Under multi-round conditions to examine the stead y-sta te rates of transcription, we observed that the fluorescent time traces exhibited an apparent lag, followed by a linear incr ease in fluor escence over time.This rate of incr ease slows over longer timescales and eventually begins to plateau (Figure 3 A, solid lines).We hypothesized that saturation of the fluorescence signal can be explained by the presence of paused and / or backtr acked polymer ases tr apped on the template (re vie wed in ( 53 )), leading to a reduction in the RNAP molecules tha t can re-initia te a t the promoter over time and an inability of active polymerases to complete transcription of a full length RNA.Consistent with this hypothesis, traces collected with E. coli RNAP in the presence and absence of the RNA cleavage factor GreB overlap at early timepoints and then di v erged, where conditions containing Gr eB r esulted in a higher fluor escent signal (Supplementary Figure S5).Based on these results, we suggest that GreB acts to increase the acti v e RNAP concentra tion a t longer times by facilita ting recycling of long paused / backtracked elongation complexes ( 54 , 55 ).As GreB had no effect in the initial linear region, we suggest that this region reports on the true stead y-sta te ra te of initiation, without interference from inacti v e elongation states occurring downstream of the promoter.In addition, these results suggest that the fluorescent-aptamer-based assay can also be used to examine the regulation of rate-limiting elongation processes at long timescales.
Based on these results, we analyzed the initial fluorescence increase to quantitate the steady-state rate.Previous work manually determined a time regime of the trace to fit with either a line or an exponential.The slope was then determined directly or through dif ferentia tion, incorpora ting an initial time offset to account for the lag-phase ( 43 ).To define the stead y-sta te (linear) regime, we designed an automated fitting protocol to reproducibly fit large amounts of data.With this strategy, there is no need to pre-determine either a time range to fit or a functional form r epr esenting the entire time course, both of which can change depending on the conditions of individual experiments.First, based on inspection of the traces to be fit, the user defines an initial time for the fitting procedure.In practice, this time should be past any lag or non-linear initial phase in the data.Then, each trace in the input data set are recursi v ely fit with a line, using a statistical weight determined by the variation between technical replicates.The length of the time frame in each round of fitting is reduced by shortening the fit time frame (i.e.decreasing the time of the final data point considered in the fit), and the R 2 value (goodness of fit) is compared to a threshold set by the user to evaluate whether the fit was adequately linear or whether a further reduction in the time frame should be attempted (Supplementary Figure S6).The code then plots the raw data and the fit for visual inspection by the user.Finally, the estima ted ra tes are outputted for subsequent analyses.

Steady-state kinetic measurements are dependent on NTP concentration
We examined the NTP concentration dependence of reaction rates to ensure that the time-dependent signal change in fluorescence follows our expectations of steady-state behavior.In addition to promoter DNA, NTPs can also be considered as a substrate in the Michaelis-Menten analysis of the transcription reaction.In this case, concentrations below those r equir ed to r each V max can lead to a decr ease in the rate of transcription, generally attributed to slower nucleotide incorporation rates and / or increases in the aborti v e fr action ( 14 , 20 ).Titr ating all four NTPs together up to 500 M on the rrnA P3 circular plasmid template, we observed a clear concentration dependence in the real-time traces (Figure 3 A, solid lines).We fit the entire data set with our variable-time fitting algorithm described above, permitting us to extract the stead y-sta te ra tes a t each NTP concentration (Figure 3 A, dotted lines).We plotted these rates as a function of NTP concentration and fit the data to a modified Michaelis-Menten equation, weighted by the exponential parameter n (Equation 2 , Materials and Methods) (Figure 3 B).The data did not fit the simpler form of the equation where n = 1 and an unconstrained fit results in an n = 2.1 ± 0.4.When n > 1, it signifies a steeper concentration dependence than expected from Michaelis-Menten equation for a single substrate binding site.Possible interpretations for this non-hyperbolic behavior can be found in the Discussion.
We performed analogous NTP titrations on the pro-moterLESS cir cular plasmid template.Compar ed to the kinetic parameters obtained with the rrnA P3 circular plasmid, we observed an ∼25-fold decrease in V max (6.5 ± 3.8 AU / min compared to 165 ± 7 AU / min) and an ∼5-fold increase in the apparent K m ( K m,app defined in Methods; 230 ± 120 M compared to 44 ± 5 M) (Figure 3 B, inset).As mentioned abov e, gi v en that the signal from the promoterLESS circular plasmid is above that of the no-aptamer control obtained a t sa tura ting NTPs (Figure 2 A), we interpret the NTP-dependent signal using the pro-moterLESS template as a measure of the non-rrnA P3 promoter deri v ed background transcription.Being ab le to measure these NTP concentration-dependent trends in signal demonstra tes tha t the assay possesses the detection sensitivity to measure low rates of full-length transcription, including those produced from sequences nominally devoid of promoters.As the promoterLESS control sets a lower bound for detecting rrnA P3 promoter-dependent transcription, it should always be included and, if need be, used as a correction.In this specific case, subtraction of the pr omoterLESS signal fr om that obtained with rrnA P3 resulted in no significant change in Michaelis-Menten parameters V max , K m,app or n (data not shown).These results indica te tha t we can confidently assign the fitted kinetic parameters to a rrnA P3-deri v ed product under these experimental conditions.

Steady-state measurements show expected dependence on both RNAP and DNA concentration
In bacterial cells there exists a large molar excess of genomic DNA relati v e to the amount of free RNAP, making the in vivo global rate of transcription largely independent of gene concentration (56)(57)(58)(59).Under these conditions, if the free RNAP concentration is well below the K m , the initial rate becomes independent of DNA concentration and proportional to the free RNAP concentration.As a result, the specific ratios of total RN AP concentration, free RN AP concentration (i.e.RNAPs that are not non-specifically bound to DNA or acti v ely performing transcription ( 60 , 61 )) and the number of possible interaction sites on the DNA will determine the reaction r ate.To illustr ate these concepts in vitro , we performed DNA titrations at multiple RNAP concentrations, using both circular plasmid (2557 bp) and linear PCR templates (250 bp) to provide different numbers of non-rrnA P3 specific interaction sites for the RNAP.
Pr evious measur ements using an aptamer-based assay with T7 RNAP illustrated that the DNA concentration dependencies of stead y-sta te ra tes followed a hyperbolic curve and could be described by Michaelis-Menten kinetics ( 39 ).We performed titrations of the rrnA P3 circular plasmid (0.1-50 nM) at two different concentrations of Mtb RNAP (20 and 100 nM) (Figure 4 A, B) and observed significantly higher rates at the higher RNAP concentration (Figure 4 C) as expected given the excess of RNAP over DNA.However, we did not observe a monotonic increase in the rates as a function of DNA concentra tion a t either RNAP concentr ation.At low nM concentr ations of the plasmid, we observed the expected increase in stead y-sta te ra te; however, as the plasmid concentration was increased, the rate exhibited a concentration-dependent decrease rather than a plateau (Figure 4 C).Of note, the peak velocity at both concentrations of RNAP occurred at a similar DN A:RN AP ratio, roughl y w hen RN AP was in 10-fold excess of plasmid DN A (Figure 4 D).These observations are consistent with a mechanism where rrnA P3 promoter-specific DNA binding dominates in conditions of limiting DNA concentration, but as the concentration of total binding sites is increased, RNAP binds to sites other than rrnA P3 mor e often, r esulting in a reduction in specific activity.
Analogous titrations done on linear PCR templates containing the rrnA P3 promoter sequence (see Supplementary Table S2 for template preparation, description, and sequence), (Supplementary Figure S7A, B) displayed a less pr evalent r eduction in stead y-sta te ra tes a t higher DNA concentrations (Supplementary Figure S7C, D).Specifically, the reduction in stead y-sta te ra te did not occur until reaching a DN A:RN AP ratio of ∼2:1 for the 20 nM RNAP condition (Supplementary Figure S7D) and no reduction was observed in the data collected with 100 nM RN AP over the DN A concentr ation r ange tested (Supple-mentary Figure S7C,D).Thus, on these short linear PCR templates, the maximal transcriptional activity occurred at a higher DN A:RN AP ratio compar ed to cir cular plasmid templates.A shift to a higher ratio of the peak velocity may be caused by a decrease in non-rrnA P3 sites.When we normalize the DNA concentration by length, the maximal activities for both linear PCR and circular plasmid templates over lay ed, suggesting that the rate decrease is due to non-rrnA P3 specific binding (Figure 4 D).Combined, these experiments demonstra te quantifica tion of the dependencies of the stead y-sta te ra tes on both RNAP and DNA concentration, as well as highlight the advantages of using low DN A concentrations w hen quantifying promoter-specific initia tion ra tes.This is especially important on plasmid templates w hich typicall y contain a higher number of possible interaction sites other than the promoter sequence under study.

Comparisons between fluorescence and gel-based approaches
Our fluor escence measur ements clearly exhibit a promoter dependence (Figure 3 B).As expected, performing analogous low throughput experiments involving incorporation of 32 P labeled UTP followed by separation via polyacrylamide gel electrophoresis (Methods) re v ealed a specific band of the expected length in accordance with the position of the termination sequence on the circular plasmid containing the rrnA P3 promoter but not on the promoter-LESS circular plasmid template (Figure 5 A).In addition, bands larger than the full-length product were observed with both templates (Supplementary Figure S8A).As these bands were of equal intensity in both the rrnA P3 and pro-moterLESS da ta, we concluded tha t their genera tion was due to a component of the plasmid (possibly other cryptic promoters found on the plasmid) other than the promoter region being studied and they are not considered further.
We next asked whether the time evolution of the promoter-specific band yielded the same kinetics as those measur ed in r eal-time via the aptamer assay.For this comparison, gel-based experiments were performed at 20 and 100 nM Mtb RNAP with 5 nM rrnA P3 circular plasmid DNA (Figure 5 B; Supplementary Figure S8B).We then normalized both sets of data using the 30-minute timepoint from the 20 nM RNAP condition and compared the time dependences of each signal (Figure 5 C).With the exception of the 100 nM RNAP 20-and 30-minute timepoints, the normalized curves overlap within error.Additionally, a comparison of the two approaches shows that the normalized stead y-sta te ra tes a t each RNAP concentra tion are consistent (Figure 5 C, inset).This analysis provides further evidence that the fluorescence signal reports on the promoterspecific activity of the system.Furthermore, it suggests that one may cautiously use the calcula ted RNA concentra tion of the specific, promoter-deri v ed band from gel quantifications to convert the arbitrary fluorescence signal into an estima ted RNA concentra tion.Fluorescence signal and RNA concentration were plotted against one another to construct a calibration curve omitting the long time points from the 100 nM RNAP condition (Figure 5   concentrations.In particular, it suggests a lower limit of detection via the aptamer assay of a pproximatel y 1 nM, consistent with previous estimates ( 41 ).The slope of this calibration was 75 ± 17 AU / nM RNA.We note this conversion factor will not be uni v ersal and will depend highly on the experimental setup and conditions (see Supplementary Discussion).In summary, results from the gel-based and fluorescence-based assays are consistent and performing both assays provides a means of converting the arbitrary fluorescence signal into absolute RNA concentration.

The spinach aptamer and DFHBI have no effect on the measured steady-state rates
We have demonstrated that use of the aptamer assay allows for the quantification of stead y-sta te ra tes of transcription.Howe v er, the two additional elements that could theoretically alter the measured rate have not been discussed: the aptamer sequence and the DFHBI dye.To examine the effect of the aptamer sequence or the presence of the dye on the rate of transcription, we performed 32 P incorporation assay measurements on circular plasmid templates containing the rrnA P3 promoter in the presence and absence of the aptamer sequence and in the presence and absence of dye.We observed roughly the same stead y-sta te ra te ± aptamer sequence (Supplementary Figure S9A, B), suggesting that the fluorescence assay can be taken to report on the rates of transcript production without effects from the aptamer.Additionally, in titrating DFHBI concentration up to 20 M (the concentration used in the fluorescence assay) in 32 P incorporation e xperiments, we observ ed no change in the amount of RNA generated 30 min after initiating the transcription reaction (Supplementary Figure S9C, D), consistent with previous measurements made with T7 RNAP ( 39 , 62 ).
We note that in the aptamer assay, we observe a change in the magnitude of the fluorescent signal when titrating dy e concentr ations in the context of the same transcription reaction (Supplementary Figure S10A).This can be explained by the finite affinity of the dye for the aptamer.As dy e concentr ation is increased, higher and higher fractions of aptamer are bound, effecti v ely increasing the gain of the signal.Howe v er, if e xperiments are al wa ys perf ormed at a fixed dye concentration, preferably higher than the K d to maximize signal (Supplementary Figure S10B), comparing fold-changes in fluorescence is a valid approach under stead y-sta te conditions (Supplementary Figure S10C, D).Care should be taken when using large concentrations of dye, as a correction for an inner filter effect may be needed ( 63 ).

High-throughput capabilities of the fluorescent assay permits concentration dependencies of individual NTPs to be measured in a single experiment
Higher concentrations of the initiating nucleotide (iNTP) than the subsequent NTPs are frequently required for promoter-specific initiation, especially on ribosomal RNA promoters where incorporation of the iNTP increases the population of open complexes at equilibrium, which can be a rate-limiting step ( 20 , 64 ).We hypothesized that when measured under stead y-sta te conditions, titra tions of the iNTP would thus yield the highest K m compared to the other NTPs on the Mtb ribosomal RNA promoter.To measure the dependence of stead y-sta te ra tes on the concentration of individual NTPs on the rrnA P3 circular plasmid template, we performed titrations of each NTP from 2.5-500 M in a background of sa tura ting concentra tions (500 M) of the other three NTPs (Figure 6 A).Each independent experiment incorporated three technical replicates across ten concentrations per individual NTP titration resulting in a total of 120 total kinetic measurements obtained in parallel, highlighting the high-throughput nature of this assay.The resulting traces were fit to extract stead y-sta te rates as a function of individual NTP concentration (Figure 6 B).When the iNTP (GTP) or the second incorporated NTP (UTP) were titrated, the Michaelis-Menten equation (Equation 1) was unable to account for the data.For these titra tions Equa tion 2 was used to account for the sigmoidal dependence.The V max for each titration was equal within error (Figure 6 A,B; Supplementary Table S3).As predicted, the data re v ealed that the transcription rate depends most strongly on the iNTP (GTP) concentration with a K m,app of 16 ± 2 M, consistent with the results from single timepoint experiments on E. coli ribosomal RNA promoters ( 64 ).As the first incorporation site of the titrated nucleotide is found further downstream within the sequence of the initially transcribed r egion, the measur ed K m,app shifted to lower concentrations (Figure 6 B; Supplementary Table S3).Notably, these trends in K m,app and V max as a function of nucleotide identity were recapitulated using short linear PCR templates containing the rrnA P3 sequence (Figure 6 C, D).Furthermore, the K m,app values were either within error or higher in all cases compared to those determined with the circular plasmid template (Figure 6 D; Supplementary Table S3), consistent with previous linear and plasmid template comparisons made on ribosomal RNA promoters ( 64 , 65 ).Even though we observed a substantially lower V max when considering the scale of arbitrary fluorescent units on linear as opposed to plasmid templates, the fact that these nucleotide-identity-dependent trends could be easily measured on either template, including the sigmoidal concentr ation dependencies, demonstr ates the utility of the assay regardless of the type of template used.

Quantification of transcription factor activity under steadystate conditions
W hen evalua ting the ef fect of a transcription factor in vitro , the most common approach is to use single timepoint  S3.
measurements to compare transcription in the presence and absence of the factor.Howe v er, by only evaluating a single timepoint for each condition there is no guarantee or confirma tion tha t the measur ements ar e r epr esentati v e of changes occurring throughout a stead y-sta te process.In contrast, the aptamer assay directly measures the effect of transcription factors in real-time and by focusing the analysis on the linear regime in an unbiased manner, ensures that factordependent changes are quantified under stead y-sta te conditions.To demonstrate the use of the assay in analysis of transcription factor effects, we turned to two well-studied Mtb transcription factors, CarD and RbpA.
Pre vious studies hav e shown that both CarD and RbpA stabilize the open complex ( 18 , 19 , 66 , 67 ), slow promoter escape ( 20 ) and activate transcription from the rrnA P3 promoter ( 66 , 68 ).In addition, these factors are known to bind the initiation complex cooperati v ely and act synergistically ( 19 , 20 , 67 ).We measured transcription in the presence of CarD, RbpA, or both at satur ating concentr ations on the linear PCR rrnA P3 template.We monitor ed fluor escence over time and fit the traces to obtain stead y-sta te ra tes (Figure 7 A).Between 2 and 5 independent experiments were used to calculate and compare the average rates at each condition.Consistent with previous work, we observed a 2.8f old, 4-f old and 9.9-fold increase in the rate of RNA production in the presence of RbpA, CarD and both factors together, respecti v ely (Figure 7 B).

Quantification of inhibitory concentrations of antibiotics
Bacterial RNAPs are the direct targets of many antibiotics (re vie wed in ( 12 , 69 )) and the search for ne w antibiotics to overcome drug resistance is ne v er-ending.This is particularly important in the battle against Mtb , the causati v e agent of tuberculosis, as multi-drug resistant strains are becoming more prevalent ( 70 ).To this end, the  de v elopment of ne w small molecule inhibitors that work at the le v el of transcription is of high inter est.Her e, we demonstra te tha t the aptamer assay can be used to quantify antibiotic-dependent inhibition of stead y-sta te ra tes.We illustrate this with the well-characterized antibiotics Rifampicin and Fidaxomicin, currently used to treat Mtb and Clostridium difficile infections by directly targeting the bacterial RNAP (71)(72)(73)(74).
Titrations of both Rifampicin and Fidaxomicin were performed using either 100 nM Mtb or E. coli RNAP with 5 nM Mtb rrnA P3 circular plasmid template (Supplementary Materials and Methods), and the kinetic traces were fit to extract the stead y-sta te ra tes.Although E. coli RNAP exhibits a lower maximal rate of transcription on this template, as inferred from the fluorescence signal in the absence of antibiotic (Supplementary Figure S11), for both the RNAPs, we observed a concentration-dependent decrease in the measured stead y-sta te ra tes upon increasing antibiotic concentrations (Figure 8 ).Fitting the data to an inhibition curve (Equation 3) permitted the calculation of half-maximal inhibitory concentrations.Consistent with pre vious wor k, the IC 50 for Rifampicin on E. coli RNAP (15 ± 2 nM) and Mtb RNAP (17 ± 2 nM) were within error (Figure 8 A) ( 66 , 75 ).In addition, the same analysis for Fidaxomicin yielded an a pproximatel y three-fold higher IC 50 on E. coli RNAP (400 ± 110 nM) than on Mtb RNAP (138 ± 92 nM) (Figure 8 B), consistent with previous reports of Mtb being more sensiti v e to the drug than E. coli ( 72 , 76 ).

High-throughput kinetic measurements of full-length transcription
The fluorescence experiments presented here are an attracti v e alternati v e technique to the standard radiolabelednucleotide incorporated gel-based a pproaches w hen used to quantita te stead y-sta te transcription kinetics, especially when used in a plate-reader format as described.Hundreds of conditions can be measured in real-time with se v eral technical replicates in a single experiment.We estimate that the throughput is on the order of a hundred times that of standard gel-based a pproaches, w here the real-time measurements facilitate a more precise and accurate determina tion and quantifica tion of the stead y-sta te regime.As the stead y-sta te ra te of full-length transcription (along with the RNA degr adation r ate) is a crucial metric when considering the flux of biological processes that use RNA as a substrate ( 1 ), use of this assay will significantly facilitate the exploration of di v erse mechanisms of gene regulation with biological relevance.
We emphasize that radioacti v e techniques are advantageous and complementary when looking to resolve individual RNA transcripts, as well as to determine the relati v e lengths and amounts of side and aborti v e products ( 77 ).In fact, the first stead y-sta te assays tha t reported on bacterial transcription were made under conditions where NTP synthesis was restricted to aborti v e transcription ( 78 , 79 ).Howe v er, as RNAP does not escape the promoter under these circumstances, these assays did not report on the overall ra te of initia tion, but ra ther the ra te-limiting step involved in binding and isomerization to the catal yticall y acti v e open complex ( 79 ).Fluorescent-labeled nucleotides that are incorporated into the RNA transcripts have also been used to monitor the stead y-sta te ra tes of aborti v e RNA generation with E. coli RNAP ( 80-83 ), r epr esenting an alternati v e to radioactivity-based approaches.However, this method cannot discriminate between aborti v e and full-length RNA products.In comparison to other techniques, the fluorescent aptamer-based assay described here is advantageous, as the observed fluorescence signal reports specifically on full-length RNA products.This is demonstrated in conditions where single NTPs are not included, resulting in the inability of RNAP to escape the promoter and a lack of fluorescence signal change (Figure 6 ).We note that the effects of aborti v e synthesis are still borne out in the kinetics, as the process of promoter escape can be rate-limiting on some promoters and affected by the initially transcribed sequence ( 84 , 85 ), which we have included in our DNA constructs.As a result, the assay permits a straightforward way to measure the rate of full-length transcript production dictated by the over all r a te of initia tion.
We have illustrated that the assay can easily quantita te dif ferences in kinetics due to promoter sequence  S11 for the corresponding real-time data, linear fits, and un-normalized rates.The normalized stead y-sta te ra tes are based on the associa ted fits using Equa tion 3 (Supplementary Figure S11E, F).
(Figure 3 ), NTP concentration (Figure 6 ), and RNAP and promoter DNA concentrations (Figure 4 ; Supplementary Figure S7).The non-monotonic rate dependencies observed when titrating DNA (Figure 4 ; Supplementary Figure S7) suggest one could use this assay to evaluate how the presence of other promoters or number of non-promoter sites compete for RNAP binding and affect the steady-states rates from a particular promoter of inter est.Furthermor e, in addition to having great utility in multi-round assays, the assay can be used in single-round conditions to evalua te ra tes of initia tion processes such as promoter escape (Supplementary Figure S4).The single-r ound appr oach has previously been well-described and used for monitoring cotranscriptional RNA folding processes ( 43 , 51 ).
We stress that meaningful quantifications of relati v e changes in flux can be made using arbitrary fluorescence units as long as buffer conditions , temperature , or any other variable that may affect aptamer folding or DFHBI binding are not varied.Howe v er, we hav e also shown the approach and necessary controls to calibrate the fluorescent signal using an independent measure of RNA concentration from identical reactions (Figure 5 ).Further thoughts on calibration and interpretation of the fluorescent signal can be found in the Supplementary Discussion.

Nucleotide-dependent kinetics
By performing titrations of NTP substrates, we have illustra ted tha t K m,app and V max parameters can be obtained with a high le v el of precision under stead y-sta te conditions (Figure 6 ).Using these data, we observed that not all titrations could be fit to a hyperbolic Michaelis-Menten model.Rather, sigmoidal trends were observed for both circular plasmid and linear PCR DNA templates in the titration of all NTPs together and GTP in the background of sa tura ting concentrations of ATP, UTP, and CTP (Figure 6 B, D; Supplementary Table S3).Initiation r equir es the first two nucleotides to form the first RNA phosphodiester bond and, depending on the conformation of the open DNA, basestacking with the +1 / +2 DNA sequences on the template strand may r epr esent a rate-limiting step ( 17 ).Our r esults are consistent with this hypothesis in that we observe that the concentration of the first two NTP substrates had a larger effect as compared to the other NTPs (Figure 6 B, D; Supplementary Table S3).For monomeric enzymes containing a single acti v e site, a sigmoidal dependence of velocity on substrate concentration has been r eferr ed to as kinetic cooperativity and was first measured with glucokinase ( 86 , 87 ).Additionally, other mechanisms have been suggested to lead to kinetic cooperativity without direct interactions between binding sites (re vie wed in ( 49)), including the existence of a slow conformational change that precedes substr ate incorpor ation, the existence of multiple enzyme classes capable of binding initiating nucleotides with dif ferent af finities, and / or a substra te-induced conformational change to the catal yticall y acti v e enzyme ( 49 , 88 ).Mechanistic explanations may include effects of the first two nucleotides on the ratio of aborti v e and producti v e synthesis or the necessity to bind two NTP substrates to synthesize the first phosphodiester bond.These models may not be m utuall y e xclusi v e and, in fact, are supported by numerous e xperimental studies e valuating the initial nucleotide incorporation process with E. coli RNAP ( 13 , 14 , 16 , 17 ).While we cannot directly comment on the specific mechanism(s) that results in this apparent positi v e cooperati vity with Mtb RNAP, the detection of this sigmoidal behavior would not hav e been possib le without the fine sampling of NTP concentrations and pr ecise measur ements of steady-state rates facilitated by the ease and throughput of the aptamer-based assay.

Future uses for high-throughput detection of basal and regulated transcription kinetics
Gi v en these fluorescent experiments were all performed in a 384 well plate, the assay lends itself naturally to questions that r equir e large data sets.We conclude by summarizing a fe w e xciting possib le directions.
Using sequence to predict transcriptional activity remains a challenge, as the context of the entire promoter must be considered when examining how sequence affects initiation kinetics ( 3 ).Using the aptamer-based assay, one  ( 44 , 84 , 89 ).Alternati v ely, one could design and measure basal and regulated transcription rates on large numbers of genomic promoter sequences.This would allow for a side-by-side comparison with genomewide, RNA-seq data ( 90 ).Use of the aptamer-based assay will greatly facilitate the ability to quantitati v ely probe the kinetics of larger genomic promoter libraries in vitro to expand our knowledge of gene regulatory mechanisms.
As we illustrated the utility of the fluorescence assay in measuring antibiotic inhibition (Figure 8 ; Supplementary Figure S11), the high-throughput screening of novel inhibitors against the bacterial transcription machinery may also benefit from this assay.Inhibitors that act at the le v el of initiation could be identified by a reduction of the initial stead y-sta te ra te, as observed for Rifampicin and Fidaxomicin (Figure 8 ; Supplementary Figure S11).In addition, inhibitors of elongation or termination that lead to stalled, template bound RNAPs may result in changes in the kinetics of the signal over longer-timescales as they e v entually reduce the number of acti v el y transcribing pol ymerases and may become roadblocks to RNAPs that are still acti v e.As a result, use of this assay under multi-round conditions may permit the identification of compounds that inhibit transcription via different mechanisms.Furthermore, because one tracks the time-dependent changes in fluorescence and not single-timepoints, compounds that exhibit fluorescent spectral properties are less likely to produce artifactual results.Compounds identified as potential hits can then easil y be anal yzed via titrations to measure the IC 50 s as described here.To de v elop structural hypotheses for the mechanism of transcription initiation inhibitors (or classes of inhibitors), titrations of NTPs or promoter DNA could be performed to determine the type of inhibition (e.g.competiti v e, non-competiti v e, or uncompetiti v e) via quantification of changes in K m,app and / or V max ( 91 ).

Figure 1 .
Figure 1.Ov ervie w of the fluorescent-aptamer-based assay.This assay requires RNAP, a DNA template containing the sequence for the Spinachmini RN A a ptamer ( 33 ), NTPs and the fluorescent dye DFHBI.Upon initiating the reaction, full-length RNAs containing the Spinach-mini aptamer capable of binding DFHBI are synthesized.The fluorescent signal change, monitored in real-time, results from the unbound to aptamerbound dye transition and is used as a reporter of full-length transcription rates.An initial lag time is observed, followed by a stead y-sta te regime where the slope of the linear phase represents the stead y-sta te ra te of fulllength RNA synthesis.

Figure 2 .
Figure 2. Real-time fluorescent signal is promoter dependent and due to multiple rounds of initiation.All experiments were performed using 100 nM Mtb RNAP, 5 nM DNA circular plasmid templates, and 500 M NTPs.( A ) Comparison of time courses of RNA synthesis from an Mtb plasmid template containing either both the rrnA P3 and aptamer sequences (black), the rrnA P3 sequence with no aptamer sequence (blue), or the 'promoterLESS' template, containing the aptamer, but lacking the rrnA P3 sequence (red).( B ) Real-time traces comparing multi-round (black) and single-round conditions (gold) on the rrnA P3 circular plasmid template.Error shading indicates the standard deviation of 3 independent experiments.

Figure 3 .
Figure 3. Quantification of the NTP-dependence of stead y-sta te ra tes for full-length multi-round tr anscription.( A ) Titr ation of the concentr ation of all NTPs with 100 nM Mtb RNAP and 5 nM rrnA P3 circular plasmid DNA results in an increase in the rate and amplitude of the fluorescent traces (solid, coloured lines).The unbiased linear fits of the early times are shown in grey dotted lines for each trace.( B ) Quantification of stead y-sta te ra tes on rrnA P3 (black) and promoterLESS (red) circular plasmid DNA templates plotted as a function of NTP concentration and fit to a modified Michaelis-Menten model (Equation2) to account for the apparent sigmoidal behaviour.
D).The best fit line (Figur e 5 D, r ed) r esulted in a negati v e y-intercept suggesting that the radioacti v e assay is more sensiti v e at low RNA

Figure 4 .
Figure 4. Stead y-sta te ra tes exhibit a biphasic DNA-concentra tion-dependence a t m ultiple RN AP concentra tions.Real-time da ta obtained a t 500 M all NTPs, titrating Mtb rrnA P3 circular plasmid DNA template (0.1-50 nM) at either (A) 20 nM or (B) 100 nM Mtb RNAP concentrations .The unbiased linear fits of the early times are shown in grey dotted lines for each trace.(C) Stead y -sta te ra tes obtained from the linear fits in (A) and (B) for 20 nM (green) and 100 nM (black) RNAP, plotted as a function of rrnA P3 circular plasmid DNA concentration.(D) Steady-state rates, normalized from zero to one based on the lowest and highest rate obtained at each RNAP concentration, plotted as a function of the normalized [DN A]:[RN AP] concentration r atios.Included titr ations of steady-state rate data are those obtained from the circular plasmids templates (open circles) and from linear PCR templates (Supplementary Figure S7, closed circles).Linear PCR DNA template concentration was normalized to that of the plasmid by dividing by a factor of ten to account for the template length (linear PCR template = 250 bp; circular plasmid template = 2557 bp).

e99 19 PAGE 10 OF 16 Figure 5 .
Figure 5.Comparison and calibration of gel-and fluorescence-based kinetics.Transcription gels showing the increase in amount of the specific, full-length 32 P-labeled transcript with time (see Supplementary Figure S8 for full gel images) obtained with (A) 100 nM Mtb RNAP and 5 nM of the promoterLESS or rrnA P3 circular plasmid templates and (B) 20 nM and 100 nM Mtb RNAP with 5 nM rrnA P3 circular plasmid template.(C) Comparison of timedependent signals from the fluorescent-aptamer-based assay (lines) and the gel assay (open circles) as a function of time for 20 nM and 100 nM RNAP.Both gel data sets were divided by the signal at 30 min obtained with the 20 nM RNAP concentration, thus normalizing that signal to a value of 1.The fluorescent data sets were then normalized using a factor determined by the ratio of the fluorescent and gel da ta a t the same timepoint.The two gel data points tha t devia te from the fluorescent time course are indicated with red asterisks.The inset shows a comparison of normalized stead y sta te ra tes from the gel (open bars) and fluorescence (filled bars) data with 20 nM (green) and 100 nM (black) RNAP concentrations.(D) Fluorescent signal plotted against RNA concentration from the data shown in (B) excluding the 20 and 30 minute time points obtained with 100 nM RNAP.A linear fit (solid red line) along with 95% confidence regions (dashed red lines) are indicated.The best fit line has a slope of 75 ± 17 AU / nM RNA and an y-intercept of -125 ± 70 AU.Error bars indicate the standard deviations from 3 to 4 independent experiments for fluorescence data and 2-3 independent experiments for gel-based data in all sub-plots.

Figure 6 .
Figure 6.Titrations of individual NTPs re v eal incorporation of the initiating nucleotide is rate limiting.(A) Real-time fluorescent traces and linear fits obtained using 100 nM Mtb RNAP, 5 nM rrnA P3 circular plasmid DNA, and individually titrating GTP, UTP, CTP, or ATP in the presence of 500 M of the other three, non-titrated NTPs.(B) Rate dependence on the concentration of individual NTPs from the data in (A) as well as the titration of all NTPs equally.Note that for clarity, only the titration data out to 200 M NTPs is shown.The y-axis was converted to RNA concentrations using the calibration presented in Figure 5 D. (C) Real-time fluorescent traces and fits as in (A), except using 25 nM of the rrnA P3 linear PCR DNA template.(D) Rate dependence on the concentration of individual NTPs from the data in (C) as well as the titration of all NTPs equally.Error bars in (C) and (D) r epr esent standard deviations of 2-4 independent experiments each with 2-3 technical replica tes.Ra te dependencies in (C) and (D) were fit to Equation 1 or 2 depending on the identity of the titrated NTP, and fitted parameters are summarized in Supplementary TableS3.

Figure 7 .
Figure 7. Factor-dependent effects on steady-state tr anscription r ates.(A) Real-time fluorescent traces collected by pre-incubating 100 nM Mtb RNAP and 500 M NTPs in the presence of sa tura ting Mtb CarD (1 M) and RbpA (2 M) (solid curves) prior to addition of 25 nM rrnA P3 linear PCR template.Raw traces are shown with associated linear fits (dotted lines).(B) Comparison of stead y-sta te ra tes (average of 2-5 independent e xperiments) re v eals the extent of transcriptional activation by RbpA (red), CarD (blue), and the combination of RbpA and CarD (purple).Fold changes in the stead y-sta te ra te relati v e to Mtb RNAP alone are indicated above each condition.

PAGE 13 OF 16 NucleicFigure 8 .
Figure 8.Quantification of antibiotic IC 50 values based on changes in observed stead y-sta te ra tes.Normalized stead y-sta te ra tes plotted as a function of (A) Rifampicin and (B) Fidaxomicin concentrations for Mtb (black) and E. coli (red) RNAP.All experiments were performed using 1 mM of each NTP and 5 nM rrnA P3 plasmid DNA.See Supplementary FigureS11for the corresponding real-time data, linear fits, and un-normalized rates.The normalized stead y-sta te ra tes are based on the associa ted fits using Equa tion 3 (Supplementary FigureS11E, F).

e99
Nucleic Acids Research, 2023, Vol.51, No. 19 PAGE 14 OF 16 could generate promoter libraries where sequence context is investigated by introducing sequence mutations in promoter r egions of inter est