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Regulation of Steady State Ribosomal Transcription in Mycobacterium tuberculosis:
Intersection of Sigma Subunits, Superhelicity, and Transcription Factors.
Ana Ruiz Manzano1,†, Drake Jensen1,†, and Eric A. Galburt1,*
1Department of Biochemistry and Molecular Biophysics,
Washington University School of Medicine, Saint Louis, MO, USA, 63108
*To whom correspondence should be addressed.
egalburt@wustl.edu, +1 314-362-4821
†These authors contributed equally
ABSTRACT
The regulation of ribosomal RNA (rRNA) is closely tied to nutrient availability, growth
phase, and global gene expression, serving as a key factor in bacterial adaptability and
pathogenicity. Mycobacterium tuberculosis (Mtb) stands out from other species with a single
ribosomal operon controlled by two promoters: rrnAP3 and rrnAP1 and a high ratio of sigma (σ)
factors to genome size. While the primary σ factor σA is known to drive ribosomal transcription,
the alternative σ factor σB has been proposed to contribute to the transcription of housekeeping
genes, including rRNA under a range of conditions. However, σB’s precise role remains unclear.
Here, we quantify steady-state rates in reconstituted transcription reactions and establish that σA-
mediated transcription from rrnAP3 dominates rRNA production by almost two orders of
magnitude with minimal contributions from σB holoenzymes and/or rrnAP1 under all conditions
tested. We measure and compare the kinetics of individual initiation steps for both holoenzymes
which, taken together with the steady-state rate measurements, lead us to a model where σB
holoenzymes exhibit slower DNA unwinding and slower holoenzyme recycling. Our data further
demonstrate that the transcription factors CarD and RbpA reverse or buffer the stimulatory effect
of negative superhelicity on σA and σB holoenzymes respectively. Lastly, we show that a major
determinant of σA’s increased activity is due to its N-terminal 205 amino acids. Taken together,
our data reveal the intricate interplay of promoter sequence, σ factor identity, DNA superhelicity,
and transcription factors in shaping transcription initiation kinetics and, by extension, the steady-
state rates of rRNA production in Mtb.
INTRODUCTION
Roughly one-fourth of the world population has been infected with Mycobacterium
tuberculosis (Mtb), the causative agent of Tuberculosis (TB) disease, which remains one of the
leading causes of death worldwide (1). While TB rates in the United States are relatively low, an
estimated 13 million individuals live with latent TB infections (2), characterized by an
asymptomatic, immune-controlled state (3). Without treatment, approximately 10% of those with
latent infection will experience reactivation, a hallmark that distinguishes Mtb from many other
infectious organisms (1). Mtb’s success lies in its ability to sense and adapt to changing
environments during infection, a process mainly controlled at the transcriptional level. This
adaptation relies on multiple regulatory mechanisms, including the activity of sigma (σ) factors,
global transcription regulators like CarD and RbpA, and changes in DNA topology.
Gene expression in bacteria is primarily regulated at transcription initiation, beginning with
the formation of the RNA polymerase (RNAP) holoenzyme, which requires the binding of the core
RNAP to a σ factor. This complex recognizes promoter DNA, initiating gene expression programs
that enable the bacterium to respond to different conditions. Mtb has 13 σ factors (4), giving it the
highest ratio of σ factors to genome size among human pathogens (5). These σ factors
orchestrate gene expression during initial infection, adaptation to stress, and the transition from
latent to active disease (6). Among them, σA is the essential housekeeping σ factor (7–9) in Mtb
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and is analogous to Escherichia coli σ70 (10, 11). Mtb σA holoenzyme is responsible for
transcribing genes required for exponential growth, including those for RNA polymerase,
ribosomes, and metabolism (9, 12). In contrast, Mtb σB, a non-essential σ factor, plays a key role
in stress responses and survival, yet also exhibits activity during exponential growth (13). Mtb σA
and σB share highly homologous C-terminal regions (~63% identity), including those subregions
involved in recognizing the -10 and -35 promoter elements (4, 14–16). Unlike E. coli, where σS
(the equivalent of σB) is tightly regulated (17–21), σA and σB holoenzymes coexist during
exponential phase in Mtb (7, 22–24). RNA-seq (9) and ChIP-seq (25) data suggest that the σB
holoenzyme regulates hundreds of genes, including rRNA and housekeeping genes, traditionally
thought to be under the exclusive control of σA (23).
Beyond σ factors, transcription in Mtb is further modulated by global regulators like CarD
and RbpA. Mtb CarD and RbpA are essential global transcription factors required under nutrient-
rich conditions and upregulated in response to stress (26–30). Both factors directly interact with
RNAP rather than binding to specific DNA sequences (27, 31, 32), and as such, they are often
considered ubiquitous members of the initiation complex (33–35). CarD was originally identified
as a regulator of rRNA synthesis (26) and, depending on the sequence of the promoter and the
resulting intiation kinetics, may act as either an activator or repressor (31, 36–39). RbpA, on the
other hand, was first linked to disulfide stress responses (40) and later found to bind both σA and
σB holoenzymes, altering transcription initiation kinetics (16, 31, 38, 41). In addition, RbpA and
CarD function cooperatively in the context of σA-dependent transcription (31, 33, 37). Given that
both regulators affect transcription kinetics rather than directly determining promoter specificity,
their combined influence may fine-tune σ factor activity.
Another critical factor influencing transcription in Mtb is DNA topology. The energetics of
DNA unwinding—modulated by the level of negative superhelicity—affects the efficiency of
transcription initiation (42). In bacteria, genomic DNA is typically maintained in an underwound
state (43, 44) to facilitate the unwinding reactions necessary for transcription (45–50).
Furthermore, genome superhelicity can shift in response to environmental and metabolic cues
(51, 52) and, while negative superhelicity generally promotes transcription initiation, its effects
can be highly context-dependent (53, 54). For instance, in E. coli, relaxed templates enhance σ38-
dependent transcription and underwound DNA preferentially support σ70-dependent transcription
(55, 56). Alternatively, in Bacillus subtilis, σA-, σB-, and alternative σ factor-dependent
transcription appears to be largely driven by negative superhelicity (57). In Mtb, it remains
unknown whether changes in DNA topology differentially impact σA- and σB-dependent
transcription.
This interplay between σ factors, transcription regulators, DNA topology, and promoter
architecture is particularly relevant for rRNA transcription, a process that governs protein
synthesis and cellular energy balance (58, 59). Since protein translation consumes nearly 80% of
a bacterium’s ATP (60, 61), regulating rRNA synthesis is critical for survival, particularly under
stress conditions. Unlike many bacteria, which possess multiple rRNA operons (62–64), Mtb has
only one, known as rrnA (65, 66), which can be transcribed from two promoters: the principal
rrnP3 and the less-characterized rrnP1 (66, 67). The extent to which these promoters function
under different regulatory contexts remains unclear, particularly regarding σ factor usage and
DNA topology. If, as recent data suggest, σB RNAP holoenzyme transcribes rRNA under specific
conditions (9, 25), this would imply an alternative mechanism for maintaining ribosome synthesis
during stress or dormancy.
To better understand how σ factor identity, promoter usage, transcription factors like CarD
and RbpA, and DNA topology collectively regulate rRNA transcription, we measured steady-state
transcription rates across hundreds of reconstituted conditions using a real-time fluorescent-
aptamer-based assay (37). This approach allows for direct comparisons of transcription dynamics
under different conditions and provides a framework for understanding how these regulatory
elements contribute to the in vivo regulation of rRNA transcription in Mtb. Our findings shed light
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on the intricate network governing rRNA transcription showing how, under all conditions and
factors tested here, the bulk of rRNA synthesis is produced by σA RNAP holoenzyme on rrnAP3.
RESULTS
Transcription on the rrnA ribosomal promoter is overwhelmingly driven by σA on the
rrnAP3 promoter.
The mycobacterial rRNA operon belongs to the rrnA family which is typically controlled by
two or more tandem promoters. For instance, faster-growing species such as M.
smegmatis possess four promoters that regulate rRNA synthesis, whereas slow-growing species
like Mtb have only two: rrnAP1 and rrnAP3 (68). This difference suggests that variations in the
utilization of rrn promoters could be a strategy for regulating rRNA production. Prior work
evaluating the Mtb rRNA operon used LacZ reporting plasmids to illustrate that rrnAP3 showed
consistently stronger signal across all growth stages tested compared to rrnAP1 (67). In addition,
while no RNA transcripts were detected for rrnAP1, the half-life of the M. bovis σA holoenzyme on
a combined rrnAP13 template was greater than the half-life on rrnAP3 alone (69).
Here, we systematically examined
the effects of σB and σA holoenzymes on
the two rrnA promoters, rrnAP1 and
rrnAP3 using a fluorescent-aptamer-
based steady state transcription assay as
previously described (37). We constructed
circular DNA templates containing either
rrnAP1 or rrnAP3 individually, or both
promoters combined, followed by an
iSpinach-D5 aptamer sequence and
measured their transcription rates under
steady-state conditions with either σB or σA
RNAP holoenzymes (see Methods). In
each reaction, 100 nM purified
holoenzyme was pre-incubated with 5 nM
rrnAP template and transcription was
initiated by the addition of 1 mM rNTPs.
Transcription was monitored in real time
through the fluorescence enhancement
produced when the small molecule
fluorophore DFHBI binds to the folded
RNA aptamer which is transcribed. All
experiments were conducted in a 384-well
plate-reader format in 10 μl reaction
volumes at 37°C. The slope of the
fluorescence signal at long times (i.e., 500
- 1800 s) was used to ensure quantitation
of the steady-state rate of transcription
without contamination from any initial burst
phases (e.g., as particularly apparent in
the σB traces, Methods). Control
experiments using either a template with
no promoter (Figure 1, dashed black lines, white bar) or core only RNAP (i.e., no σ factor) (Figure
1A,C, dotted black lines) demonstrate that the signals are promoter and holoenzyme specific.
The data show that σA holoenzyme exhibits a 20-fold higher steady-state rate of transcription
on rrnAP3 compared to rrnAP1 (Figure 1A,B). In the case of the σB holoenzyme, this preference
Figure 1: Quantification of steady state-rates on rrnAP1, rrnAP3,
rrnAP1AP3, and promoterless circular plasmid templates initiated
with rNTPs for σA (top) and σ
B (bottom) holoenzymes. (A,C)
Comparison of real-time fluorescent signal time courses of RNA
synthesis. Template without promoter is shown with a black line.
Signal from rrnAP3 with core RNAP (i.e., no σ
factor) is depicted
with a dotted line. Shaded areas indicate the standard error of the
mean of 5 experiments. (B,D) Quantification of steady state rates
using linear fits between 500 - 1800 s. Template without promoter is
shown in grey. Data are plotted as mean ± SEM of five independent
replicates (each one performed in three technical replicates). P-
values (paired t-test) are indicated as follows: not significant (–) and
less than 0.05 (*). AP3 containing templates are significant over no
promoter with P-values of 8
´
10-7 and 2
´
10-3 for σA and σ
B
holoenzymes respectively.
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for rrnAP3 is also observed. rrnAP1 transcription by σB is not statistically different than the
template without promoter, while rates from rrnAP3 are twice as high. More significantly, overall
transcription rates from σB are over an order of magnitude lower than with the σA holoenzyme
(Figure 1C,D).
We also asked whether there may be cooperative effects on the steady-state rate of
transcription due to the presence of both promoters, rrnAP1 and rrnAP3, in tandem. Transcription
rates for σA holoenzymes on the rrnAP1P3 construct do not differ significantly from the rates
observed for rrnAP3 alone (Figure 1A,B), and transcription rate averages from the σB
holoenzyme show a small reduction, although the P value indicates no significance (P=0.083)
(Figure 1C,D). This result indicates that the rrnAP1 promoter contributes minimally, if at all, to
overall transcription under these conditions, while rrnAP3 is predominantly transcribed by the σA
holoenzyme.
σA steady-state transcription rate is two orders of magnitude faster than σB on rrnAP3 with
similar σ-concentration dependencies.
Sigma-factor concentration and availability varies subject to cell growth phases and
environmental signals (4, 15, 22, 23, 70–72), causing the concentration of corresponding
holoenzymes to change according to the affinity of each σ to core RNAP. To inform on the
activities of σA and σB holoenzymes on ribosomal transcription, we measured the concentration
dependence of both σ factors by titrating untagged σ factors in the multiround transcription assay.
Each reaction contained 100 nM core RNAP, while σ factors concentrations varied between 25
and 1400 nM. Transcription was initiated by adding 5 nM of the rrnAP3 circular plasmid
template.Due to the initiation of the reaction via the addition of DNA instead of NTPs as above,
traces lacked a burst phase (Supplementary Figure 1) and were linearly fit to extract steady-
state rates. Fits of the titrations with hyperbolic curves (V = Vmax([σ]/(Km+[σ])) allowed us to
quantitate the concentration at which half activity is reached (Km) and the maximal velocity (Vmax).
For σA, the Km is 192 ± 50 nM, while for σB, the Km is 177 ± 46 nM (Figure 2A). Thus, despite the
two-orders of magnitude difference in the maximum velocity at saturating σ factor concentrations
(92.5 ± 7.3 and 1.1 ± 0.1 AU/min respectively), σA and σB exhibit similar concentration
dependencies (Figure 2A).
The effects of CarD and RbpA on rrnAP3 transcription depend on σ factor identity and are
minor compared to the difference between σA and σB basal levels.
Given the stark decrease in Vmax with σB compared to σA holoenzymes on rrnAP3 (Figure
2A), we wondered whether the presence of either or both CarD and RbpA may specifically
stimulate σB holoenzymes to the level observed of σA, or if they merely fine-tune the transcriptional
output of each. CarD is recruited to initiation complexes in a σ-independent mechanism through
interaction with the core RNAP (73). In contrast, RbpA interacts both with core RNAP and with
the non-conserved regions of σA and σB (16, 33, 35, 41, 74). Given the global nature of these
factors (27, 75) their regulation as a function of bacterial growth (26, 76, 77), and their importance
in the response to environmental stresses (26, 77, 78), one might hypothesize that these factors
are especially important for the regulation of the steady-state rate of ribosomal transcription. To
address how transcriptional activity for each holoenzyme varied in the presence of CarD and
RbpA, we measured the σ factor concentration-dependence of steady-state transcription on
plasmids in the presence of CarD (1 µM), RbpA (2 µM) or both (Figure 2). For each σ factor
titration, we fit the data to a hyperbolic curve and extracted estimates of the concentration
dependence (Km) and the maximum velocity (Vmax) (Figure 2B,C). We then calculated the ratio of
these values to those obtained for each holoenzyme in the absence of factors (Supplemental
Table 1).
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Both factors alone and together repress σA-dependent transcription. Specifically, RpbA
reduces Vmax by 30%, while CarD reduces Vmax by 70% (Figure 2B,E). In contrast, in the case of
σB, the factors led to an increase in Vmax by up to 40-60% depending on the factor (Figure 2C,E).
Thus, with respect to steady-state transcription on circular plasmid templates, CarD and RbpA
have opposite effects on σA and σB driven transcription. Furthermore, these effects are small
relative to the overall difference between the basal levels of transcription exhibited by each
holoenzyme alone (Figure 2D). Analysis of the Km values indicates that both factors alone and
together decrease the σA-dependence. Specifically, RbpA decreases Km by 80%, while CarD
reduces Km by 150% (Figure 2B,F). In contrast, in the case of σB, the factors have an distinct
effect. Specifically, RbpA increases Km by 30% while CarD did not significantly affect it (Figure
2C,F).
DNA unwinding rate and holoenzyme recycling kinetics limit σB transcriptional output.
In an effort to determine the underlying features that bring about the large changes in Vmax
between the two holoenzymes, we measured aspects of initiation kinetics for both σA and σB
holoenzymes using linear rrnAP3 templates labeled with a Cy3 on the non-template thymine base
at position +2 in a stopped-flow assay (31, 36, 79). In the assay, promoter binding and subsequent
open complex formation leads to an increase in fluorescence intensity, whereas removal of RNAP
from the promoter either via dissociation or promoter escape leads to a decrease.
We first looked at the approach to equilibrium in the forward direction by mixing DNA and
RNAP in the absence of NTPs and tracking the fluorescence increase (Figure 3A). Based on the
Figure 2: The influence of CarD and RbpA on σA and σB driven transcription. (A) Steady-state transcription rates as a function
of σ factor concentration fit to a hyperbolic curve (σA solid line, and σB dotted line). Table insert shows the fit parameters. (B)
Titration of σA with no factors (black), CarD (blue), RbpA (red), and both factors (purple). (C) Titration of σB with no factors
(black), CarD (blue), RbpA (red), and both factors (purple). (D) A comparison of the titrations in
B
and C on a logarithmic scale.
Error bars represent standard error of the mean for each measurement and solid lines represent fits to a hyperbolic curve. (E) The
percent increase in Vma x for σA (gray) and σB (orange) holoenyzmes calculated from the data in (B) and (C) is plotted for each
factor individually and combined. (F) The percent decrease in Km for σA (gray) and σB (orange) holoenzymes calculated from the
data in (B) and (C) is plotted for each factor individually and combined.
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time to reach half-maximal signal, σA holoenzymes equilibrate three times as fast as σB (t1/2s of
31 s and 95 s respectively). Since the equilibration rate reports on the sum of the forward and
reverse rates in a two-state system, this observation suggests that either DNA unwinding, DNA
bubble collapse, or both are faster for the σA holoenzyme. Measurements of the dissociation of
these complexes by challenging with unlabeled DNA templates provided evidence for an early
phase of σB dissociation that is ~1.5x faster than that of σA (Figure 3B). Taken together, these
observations suggest an ~4.5x slower rate of DNA opening catalyzed by σB.
We also noted that the σB holoenzyme saturates at a ~25% higher fluorescence fold-
change (1.85 AU vs. 1.5 AU) suggesting that, under these conditions, σB eventually forms more
open complexes (Figure 3A). This is likely consistent with the presence of an additional and
extremely long kinetic phase (t1/2 of ~5 hrs) unique to σB in the dissociation experiments (Figure
3B,C). The multi-phasic dissociation further suggests that σB populates multiple distinct open
complex conformations.
Given the nearly two-orders of magnitude in reduction of the steady-state rate for the σB
holoenzyme, we reasoned that in addition to the ~4.5x slower DNA unwinding, perhaps promoter
escape kinetics would also be reduced. Experiments in the presence of NTPs show that escape
for both σ holoenzymes is more rapid than the equilibration of open complex, consistent with the
idea that ribosomal promoters are rate limited at DNA opening and open complex formation rather
than escape (31, 73). In addition, σB holoenzyme exhibits a similar rate of decrease in
fluorescence compared to σA (Figure 4A), suggesting that promoter escape does not contribute
to the relative deficiency of σB-dependent steady-state transcription. Alternatively, we wondered
whether the observed NTP-dependent decrease in fluorescence for σB holoenzyme in this assay
could be partially explained via processes that do not lead to the production of a full-length RNA
instead of bona fide promoter escape. This would reduce the probability of producing a transcript
for each open complex formed but would still show a fast decay rate. However, single-round
aptamer experiments revealed similar total amounts of transcript produced suggesting that σA and
σB open complexes have a comparable probability to produce transcript (Figure 4B).
By the process of elimination, these data suggest that in addition to slower open complex
formation (Figure 3A), the differences in multi-round steady-state transcription must be
significantly affected by aspects unique to the multi-round reaction. More specifically, the rates of
holoenzyme recycling (i.e., the rate of σ-core interactions needed to regenerate a competent
holoenzyme after transcription) may be a possibility.
Figure 3: Open complex formation and decay kinetics for σA and σB holoenzymes on the rrnAP3 promoter: In all panels, σA
(black) and
σ
B (orange) holoenzymes are shown. Shaded regions represent the standard deviations from multiple experiments. (A)
Fluorescent fold-change over background as a function of time showing the approach to open complex equilibrium after the
addition of holoenzyme via stopped-flow. (B) Open complex decay measured via stopped-flow showing the early phases of
dissociation. (C) Open complex decay measured via plate-reader showing the longer timescale phase unique to
σB holoenzyme.
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CarD and RbpA produce similar changes in the kinetics of open complex equilibration,
dissociation, and promoter escape in the context of σA and σB holoenzymes.
Above, we showed that σA and σB holoenzymes result in different open complex
equilibration and dissociation kinetics (Figure 3). We also showed that, under the conditions
tested, on circular plasmid templates CarD and RbpA repress steady-state transcription by σA and
activate steady-state transcription driven by σB holoenzymes (Figure 2). From theoretical (31, 80,
81) and experimental (39, 82, 83) perspectives exploring how different basal kinetics can result
in differential CarD and RbpA-based transcriptional regulation, we hypothesized that CarD and
RbpA modulate the kinetics of σA and σB holoenzymes in a similar way even though they result in
diametrical effects on steady-state rate (Figure 2). If this hypothesis is true, each factor would be
predicted to accelerate the equilibration of open complexes and slow promoter escape (31)
regardless of σ factor context. This prediction is based on the following prior work: i) on Mtb
rrnAP3 with σA holoenzymes, CarD and RbpA increase the rate of DNA opening (31, 33, 38) and
slow escape kinetics (31), and ii) on Mtb sigAP with σB holoenzymes, RbpA facilitates open
complex formation (16, 35, 83). Alternatively, we wondered if, as has been suggested for RbpA
(16, 41), there are σ-dependent differences in the activity of each transcription factor. If this picture
were true, one might expect that CarD and RbpA would produce different kinetic changes on the
same step of intiation depending on σ context.
Figure 4: Promoter escape and single round kinetics for σA and σB holoenzymes on the rrnAP3 promoter: In all panels, σA
(black) and
σ
B (orange) holoenzymes are shown. Shaded regions represent the standard deviations from multiple experiments. (A)
Normalized fluorescence as a function of time showing the decay upon mixing pre-formed open complexes with DNA competitor
and rNTPs via stopped-flow. (B) Fluorescent signal as a function of time for single-round aptamer assays in the presence of DNA
competitor. Both traces were well fit by a single exponential. Controls where DNA competitor was added prior to mixing with
NTPs (dotted lines) confirm single-round conditions.
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To test these two possibilites, we performed our kinetic stopped flow assays for each σ
factor-containing holoenzyme in the presence of CarD and RbpA individually and combined
(Figure 5). In each of the two kinetic assays, we observed that the factors changed the kinetics
in the same direction. Regardless of σ factor identity, CarD and RbpA accelerated open complex
equilibrium (Figure 5A,B) and slowed promoter escape (Figure 5C,D) and open complex
dissociation (Supplementary Figure 2). This result suggests that the distinct effects of a
particular factor on steady-state transcription rate are dictated by the different basal kinetics of σA
and σB and not by distinct molecular mechanisms. That said, while the magnitude of change in
the kinetics due to CarD is similar in the two σ factor contexts (Supplementary Figure 3A), RbpA
exhibits a quantitatively stronger effect on the kinetics of σB (Supplementary Figure 3B).
Negative superhelicity stimulates σA
and σB driven rrnAP3 transcription equally.
Knowing that DNA topology dramatically affects the energy landscape of DNA unwinding
(42), we next asked how steady-state transcription driven by each σ factor-containing
holoenzyme, and its transcription factor-dependence, varies with templates exhibiting differing
levels of superhelical densities. To investigate the influence of DNA topology on transcription
Figure 5: Dependence of forward and dissociation/escape kinetics on CarD and RbpA for σA and σB holoenzymes on the rrnAP3
promoter: In all panels, no factor (black) traces were collected under conditions where open complex was saturated and are
compared to those in the presence of CarD (blue), RbpA (red), and both factors together (purple). (A,B) Open complex
equilibration kinetics for σA and
σB holoenzymes respectively. (C,D) promoter escape kinetics for σA and σB holoenzymes
respectively. Note that the increase in fluorescence starting at ~ 100 ms before the fluorescence decay reports on initial transcribing
intermediates (see (31)) and is not discussed here further.
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rates, we utilized a set of five rrnAP3 DNA templates (Supplementary Figure 4 and Methods):
supercoiled (miniprepped and DNA gyrase treated), relaxed (Topoisomerase I treated), nicked
(Nt.BsmAI treated), and linearized (ScaI treated). Gyrase treated templates showed comparable
rates to the miniprepped DNA and are not discussed further (Supplementary Figure 5).
We first measured basal rates with either σA or σB holoenzymes in each topology. In both
holoenzymes, the supercoiled template stimulated steady-state transcription by 2-3-fold
compared to nicked and linear templates with the relaxed template displaying an intermediate
rate (Figure 6, gray). Overall, the ratios between σA and σB transcription rates are relatively
constant across the different topologies (Supplementary Table 3, no factor columns).
CarD’s and RbpA’s effects on steady-state rate differ with respect to topology and σ factor
identity
CarD and RbpA reduce or invert the topology dependence of σA transcription on rrnAP3:
In the presence of CarD, and agreeing with previous observations (39), the rate of σA-
dependent transcription on the superhelical template is slightly reduced while that of relaxed,
nicked, and linear are significantly enhanced (Figure 6A, grey vs. blue). In fact, the magnitudes
of the changes are such that the topology dependence is completely reversed in the presence of
CarD. RbpA also slightly reduced the transcription rate on the supercoiled template and
stimulated transcription on the other templates such that any topology dependence was nearly
completely removed (Figure 6A, grey vs. red). In the presence of both transcription factors, the
reversed topology dependence driven by CarD is again visible (Figure 6A, purple). The same
data are grouped in terms of topology for a comparison of the effect of each transcription factor
on a given template (Supplementary Figure 6A). Here one can more clearly see that CarD
increases rates of transcription more than RbpA in the context of σA. The fold changes relative to
no factors and relative to the supercoiled templates can be found in Supplementary Tables 2
and 3.
RbpA activates transcription across all topologies in the context of σB:
CarD has a limited effect on the transcription rates of σB holoenzyme (Figure 6B, gray vs.
blue and Supplementary Figure 6B), consistent with limited effects on open complex
equilibration and promoter escape (Figure 5). In contrast, RbpA significantly stimulates
transcription on all topologies (Figure 6B, gray vs. red and Supplementary Figure 6B), also
Figure 6: The dependence of steady-state transcription on topology and transcription factor for σA (A) and σB (B). Factor conditions
are indicated on the x-axis and topology is indicated by different shades. Error bars indicate standard error of the mean from at least
6 traces in each condition collected across multiple days, each with triplicate technical replicates. P-values (paired t-test) from the
supercoiled template are indicated by asterisks as follows: less than 0.05 (*), less than 0.005 (**), less than 0.0005 (***).
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consistent with displaying larger effects than CarD on individual initiation steps (Figure 5). In the
presence of both transcription factors, the effect is similar to that of RbpA alone (Figure 6B, purple
vs. red) As such, the preference for supercoiled templates is conserved across all conditions.
That said, the stimulator effect of RbpA is greater on the nicked and linear templates compared
to the supercoiled template (2x vs. 4x) and less so on the relaxed template (1.5x)
(Supplementary Figure 6B). Calculated fold changes relative to no factors and relative to the
supercoiled templates can be found in Supplementary Tables 2 and 3.
Differences in steady-state transcription rate are linked to the N-terminal extension of σA.
Given the stark disparity in rRNA transcription rates between σB and σA holoenzymes in
all conditions tested here, we considered potential structural determinants behind this difference.
The most notable distinction between the two σ factors is the presence of a 205 residue N-
terminal extension in σA that is almost certainly intrinsically disordered (74). Although the exact
mechanism by which this feature modulates σA function remains unknown, single timepoint gel-
based experiments on linear rrnAP3 templates suggest that, while deleting the first 179 residues
of the N-terminal extension may or may not affect promoter binding, transcriptional output is
reduced four-fold relative to full-length σA (9). Additionally, this truncation failed to complement an
in vivo model suggesting an essential functional role (9). Thus, we tested whether the N-terminal
extension in σA accounts for the observed differences between σ factors in the steady-state rates
of rrnAP3-driven transcription.
Using a mutant version of σA (σAΔ205) in which the first 205 amino acids were deleted, we
performed steady-state transcription reactions as a function of σ concentration. Each reaction
contained 100 nM core RNAP while the concentration of the mutant σ factor was titrated from 25
to 1100 nM (Figure 7A). The data show that σAΔ205 has a Vmax two orders of magnitude lower than
wildtype σA and similar to σB (Figure 7A,B). Thus, the N-terminal extension directly contributes to
the high steady-state rates observed with σA. We also tested the effect of transcription factors
CarD and RbpA on σAΔ205. These results suggest that the σA N-terminal extension also dictates
the factor-dependent effects (Figure 7B, Supplementary Table 1).
Figure 7: The effecct of the N-terminal extension of σA. (A) Steady state transcription rates as a function of σ
factor concentration
for σA (black), σB (orange), and σA
D
205 (dashed gray line). Error bars represent standard error of the mean and lines represent fits
to hyperbolic curves. (B) Percent changes in Vmax for each σ factor (σA:dark grey, σB:orange, σA
D
205:light gray with hatchmarks)
in the presence of each transcription factor.
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DISCUSSION
The expression of ribosomal rRNA genes is tightly regulated in response to environmental
signals. Both the housekeeping sigma factor A (σA) and the alternative sigma factor B (σB) are
expressed at roughly equal levels during exponential growth (23) and are associated with similar
promoters, including rRNA promoters, as revealed by ChIP-seq (25). While the σB regulon
overlaps significantly with that of σA, and it is classified as a secondary σ factor that may
complement σA function during growth (9, 25, 84), it is not essential and cannot rescue a σA
knockout (9). Notably during hypoxia or stationary phase, conditions that contribute to the
transition of Mtb into a non-replicative, dormant state, σA expression decreases 3-fold while σB
levels increase 2.5-fold (23). This coincides with a reduction in ribosomal synthesis (85), albeit to
a lesser extent compared to other bacteria (86). Here, we investigated the degree to which rRNA
transcription by σB holoenzymes contribute to the regulation of rRNA synthesis.
Ribosomal transcription output is dominated by σA holoenzymes from the rrnAP3 promoter.
Our steady-state measurements show that rrnAP3 exhibits dramatically higher output than
rrnAP1 and that the presence of both promoters does not significantly alter the rate of transcription
(Figure 1). In addition, σB holoenzymes exhibit a notably lower output compared to σA
holoenzymes across all three promoter constructs (rrnAP1, rrnAP1P3, and rrnAP3) (Figure 1).
However, these initial measurements only included the basal transcription machinery and lacked
CarD and RbpA, two global transcription factors that are often considered to be de facto members
of the complex as they are each recruited directly to initiation complexes by the RNAP
holoenzyme itself (33, 73). We considered the possibility that the presence of these transcription
factors may dramatically change the picture of how much σB contributes to ribosomal
transcriptional output. However, while each transcription factor clearly regulates steady-state
transcription driven by each holoenzyme, factor-dependent rate changes are small relative to the
nearly two orders of magnitude difference between the rates of σA and σB (Figure 2).
Taken together, our results suggest that despite observations of σB holoenzymes at
ribosomal promoters (25) and the downregulation of σA under stress and in stationary phase (23),
rRNA transcription is likely to be primarily driven by σA holoenzymes at the rrnAP3 promoter. This
result underscores the central role of σA and provides constraints when considering the role of σB
in maintaining ribosomal transcription. Yet-to-be-discovered regulatory mechanisms enacted
under specific growth conditions in specific environments in vivo may change this picture.
Mechanisms underlying differential ribosomal transcription of σA and σB holoenzymes.
Our kinetic measurements revealed that σB holoenzymes exhibit distinct initiation kinetics
in that they have a slower rate of open complex equilibration, a very long-lived open complex in
the absence of NTPs, and a similar promoter escape rate compared to σA holoenzymes (Figures
3,4). However, these kinetic and structural studies appear to represent only part of the story with
respect to large differences in steady-state rates. Since each holoenzyme exhibits similar single-
round kinetics (Figure 4B), we propose that differences in the rate of holoenzyme recycling (i.e.,
the re-binding of σ and core after a round of transcription) may impact the differences in steady-
state rates (discussed further in the section below devoted to the role of the N-terminal extension).
(87–89)(89)
We also note work showing that N terminally tagged σB holoenzymes form higher order
oligomers that may be inhibitory to transcription (35). These complexes begin to form at
concentrations of holoenzyme used here. However as they are dissolved by RbpA and we show
that the presence of RbpA does not equalize the rates between σA and σB holoenzymes (Figure
2D), the effect of these oligomers is likely subtle. Taken together, it appears that multiple
mechanisms combine to determine the relative rates of ribosomal transcription by the different
holoenzymes.
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CarD and RbpA have opposite effects on the rate of transcription of σA and σB holoenzymes on
negatively supercoiled DNA.
In the context of superhelical templates, the presence of CarD and RbpA exert opposite
effects on σA and σB driven transcription. Specifically, they repress rrnAP3 transcription mediated
by σA while activating transcription mediated by σB (Figure 2). These opposing effects on the
maximal steady-state rate are paralleled by changes in the concentration dependence of each σ
factor: CarD and RbpA reduce the Km for σA, consistent with previous studies with RbpA (41),
while increase it for σB, with an approximate two-fold difference in each case (Figure 2E,F and
Supplementary Table 1). To explore the mechanistic basis of these trends, we considered two
potential explanations. The first is that a transcription factor can lead to different regulatory
outcomes depending on the basal kinetics of a given promoter/holoenzyme system (31, 80). The
second is that a transcription factors exhibits σ factor specificity, altering initiation kinetics in a σ-
dependent manner to either repress or activate transcription (16, 41).
To distinguish between these models, we examined the effects of the CarD and RbpA on
three key aspects of initiation kinetics using a fluorescence stopped-flow assay as previously
described (31, 38, 87). Our results confirmed that CarD and RbpA induce qualitatively similar
changes in the kinetics of open complex equilibration, open complex dissociation, and promoter
escape, regardless of σ factor identity (Figures 3, 4, Supplementary Figure 2). However, within
this context, RbpA shows a quantitatively enhanced ability to modulate these kinetics in the
presence of σB compared to σA (Supplemental Figure 3). The ability of RbpA to dissolve inactive
σB holoenzyme octamers could contribute to this specificity (35).
Thus, we are left with a mixture of the two models. Overall, the direction of the regulatory
outcome arises from a shared transcription factor mechanism modulated by the basal initiation
kinetics of each promoter/holoenzyme system. Quantitatively, there also appears to be molecular
specificity, with RbpA showing a preference for enhancing σB holoenzyme activity. Together,
these results point to a complex interplay of basal kinetics and σ-specific interactions in
determining transcriptional regulation by CarD and RbpA.
Negative superhelicity stimulates ribosomal transcription by both σA and σB holoenzymes.
Negatively supercoiled DNA is maintained during growth by a balance between ATP-
dependent DNA gyrase and other topoisomerases (88). As such, genome superhelicity is also
influenced by cellular energy stores (i.e., ATP/ADP ratios). For example, during stationary phase,
when energy becomes limited and ATP levels drop, reduced gyrase activity leads to decreased
superhelicity (89). Conversely, environmental stresses like osmotic or cold stress have been
shown to increase DNA supercoiling (hypercoiling) in E. coli and B. subtilis, which can activate
transcription (88). Similarly, in Mtb, superhelicity has been implicated in the regulation of virulence
genes, such as virR and sodC, through the NapA nucleoid-associated protein (90).
Given the influence of supercoiling on transcription, we investigated how superhelical
density affects transcription driven by each σ factor. Consistent with early in vitro studies using
isolated M. smegmatis RNAP (91), we observed that both σA and σB driven transcription is strongly
dependent on the superhelical state of the DNA template (Figure 6). Specifically, transcriptional
output by both σ factor-containing holoenzymes is reduced by approximately 50% when
comparing supercoiled plasmid templates to linear templates. As negative superhelicity promotes
DNA unwinding (45–49), these observations are also consistent with a model where rrnAP3 is
rate limited in part by DNA opening (Figures 3A,4A, (31)).
CarD and RbpA buffer the dependency of σA holoenzymes on superhelicity.
To understand how the intersection of superhelicity and transcription factors influence
rRNA transcription, we examined the effects of CarD and RbpA across different σ factor and
superhelical contexts. Remarkably, we found that each factor individually, or both factors
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combined, generated a flip in the dependency of σA holoenzymes on superhelicity. Specifically,
in the presence of both factors, steady-state transcriptional output is stimulated up to two-fold on
relaxed, nicked, and linear templates compared to supercoiled plasmid templates (Figure 6). This
result is consistent with prior work illustrating that with the σA holoenzyme, both CarD and RbpA
activate rrnAP3 transcription on linear templates (37, 39, 41, 92), whereas CarD represses
transcription on supercoiled plasmid templates (39). This topological dependence on CarD activity
may also explain why in vivo, CarD depletion in M. smegmatis leads to increased rRNA levels
(26), assuming CarD directly represses rRNA transctiption on the supercoiled genome. As both
DNA topology and promoter sequence may moduulate basal transcription initiation kinetics, our
observations are consistent with prior work illustrating how changes in promoter sequence can
lead to differential regulation by Mtb CarD and RbpA (39, 83) and Rhodobacter sphaeroides CarD
(82, 93).
In the case of σB holoenzyme, a similar effect is observed, where CarD and RbpA change
the relative output. However in this instance, they reduce the disparity between linear and
supercoiled template outputs, bringing them to roughly equal levels (Figure 6). This behavior
suggests that CarD and RbpA help buffer reductions in rRNA transcription during stationary
phase, when superhelicity and σA levels are lower (23, 41, 88, 94).
On a practical experimental level, we emphasize the need to carefully control for DNA
template topology in in vitro studies. Differences in topology, whether due to the use of linear
versus circular templates or to the degradation of superhelicity via nicks in plasmids, can
significantly affect the results of transcriptional measurements and the inferred biological
mechanisms of gene regulation.
The N-terminal extension of σA is a major regulatory determinant of the steady-state transcription
rate differences between σA and σB and the effects of CarD and RbpA.
Finally, to explore potential structural determinants of the observed differences between
σA and σB, we examined the unique N-terminal domain extension in σA. Structural studies have
suggested that the N-terminal extension of σA, which σB lacks, likely plays a role in open complex
formation/stabilization due to its position in relation to the downstream DNA channel (34, 35, 95,
96). In Mtb, this region is predicted to be entirely disordered (74), while the non-homologous N-
terminal domain in E. coli σ70, termed region 1.1, is partly structured and conformationally dynamic
(97, 98). For instance, in its apo-form, σ701.1 prevents DNA binding by occluding the DNA binding
domains in a compact structure (99–101). While bound to core RNAP in a more expanded
conformation, σ701.1 occupies the active site cleft where DNA gets loaded and must be displaced
upon DNA unwinding (97, 98, 102), likely resulting in the observed changes in unwinding kinetics
(103) and complex stability (104, 105). Given the lack of sequence and structural conservation
between the Mtb σA N-terminal extension and E. coli σ 701.1 (74), the function of the σA N-terminal
extension remains unknown. As our kinetic measurements suggest that part of the difference in
steady-state rates may be due to differences in holoenzyme recycling rates, it is possible that the
N-terminal extension also affects this process.
Under the conditions explored in this paper, truncation of the first 205 amino acid residues
of the σA N-terminal extension (σAΔ205) generates steady-state rates of transcription comparable
to those of σB on plasmid templates (Figure 7A). As such, we conclude that the N-terminal
extension is a major determinant of the increased rate of transcription catalyzed by σA
holoenzymes relative to σB. Interestingly, while large changes in Vmax were observed (Figure 7B),
the EC50 of σAΔ205 was not significantly different from the EC50 of either σB or σA, suggesting a
similar affinity of each holoenzyme to promoter DNA (Supplementary Table 1). It is worth noting
that the regulatory effect of CarD and RbpA is greatly influenced by the N-terminal extension,
leading to repression of transcription when present (σA) and activation of transcription when
absent (σAΔ205), analogous to the effects observed with σB (Figure 7B). As a result, CarD and
RbpA can be added to the list of other general bacterial transcription factors, like E. coli DksA
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and TraR (102, 104, 106), and bacteriophage proteins (107, 108) whose activity is impacted by
the N-terminal region of σfactors.
Given the dramatic effect of the N-terminal extension on basal and regulated kinetics
measured here, we stress that σ factor comparisons made throughout this work utilized untagged
proteins to avoid potential artefacts of non-native sequences. Clearly, the N-terminus has the
potential to alter the complex energetics and kinetics of interactions between σ factors, the RNAP
core, and the DNA template.
It is tempting to hypothesize about the regulatory advantage provided by the N-terminal
extension into Mtb’s ability to balance transcriptional efficiency with adaptability across different
environmental contexts. While the exact molecular mechanisms remain to be elucidated,
possibilities include modulation of interactions with core RNAP subunits and sequence-specific
lineage insertions (74, 109), or other regulatory proteins (110, 111). Future studies will be needed
to determine more specific mechanisms underlying σA’s enhanced output to offer insights into the
N-terminal extension’s role in the biology of Mtb.
In conclusion, ribosomal transcription in Mtb is primarily driven by σA, with σB playing a
more limited role under the conditions studied. CarD and RbpA reverse the dependency of σA
holoenzyme on superhelicity (and buffer that of σB holoenzyme) and have opposing effects on the
transcription rates of σA and σB holoenzymes on negatively supercoiled DNA. These findings
highlight the central role of σA in meeting Mtb’s ribosomal demands and underscore the
importance of integrating direct measurements of steady-state transcription rates with techniques
like ChIP-seq and RNA-seq for a more comprehensive understanding of the regulation of gene
transcription. Further investigation into the interplay between σ factors, DNA topology, and
regulatory proteins will provide critical insights into Mtb's adaptability and survival strategies.
MATERIALS AND METHODS
Preparation of DNA constructs
For all final DNA construct sequences used in this work, see Supplementary Table 4.
Circular plasmid templates, 2557 base-pair (bp) in length, were ordered from Twist Bioscience
(San Francisco, CA) and represent an updated version of previously published constructs (37).
Sequences contain, in order, the Mtb tuf terminator (112), the Mtb ribosomal RNA promoter
(rrnAP3), the iSpinach D5 aptamer (113), and the E. coli rrnBP1 T1 terminator (114, 115).
For the preparation of topologically different DNAs, the plasmid purified from the Qiagen
Maxi Prep Kit (12963) was more than 90% supercoiled, estimated by the relative abundance of
the bands on the agarose gel (Supplementary Figure 4). For additional negative supercoiling
and removal of concatemers, the plasmid was treated with DNA Gyrase (TopoGEN, TG2000G-
1). Additionally, topologically closed but relaxed plasmids were prepared by incubation with
Topoisomerase I (NEB, M0301S). Nicked plasmid DNA was prepared by incubation with
restriction enzyme Nt.BsmAI (NEB, R0121S) that made two cuts in the plasmid upstream the
promoter region. To linearize the plasmid, the restriction endonuclease enzyme ScaI (NEB,
R3122S) was used. In all cases, a total volume of 200 μl, 20 μl assay rCutSmart buffer (NEB) 150
μl RNase-free water, 10 μl enzyme and 5 μl of plasmid DNA (c = 2000 ng/μl) were combined in a
reaction tube and incubated for 1 h at 37°C. Reactions were cleaned with a PCR clean-up kit
(Qiagen, 28104), and resuspended to 50 nM in transcription buffer (see below). Concentrations
were calculated using the molecular weight based on sequence.
A Linear 150 bp template containing the Mtb rrnAP3 promoter labelled with Cy3-NHS
(Lumiprobe Corporation, 11020) attached to a C6-amine modified thymine on the +2 position of
the non-template strand was used for plate-reader dissociation and stopped-flow assays. A biotin
molecule was added to the 5’-end of the non-template strand attached via a standard C6 spacer
(IDT). For promoter preparation and labeling protocols, see (36).
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Preparation of recombinant proteins
A summary of all expression constructs used can be found on the Supplementary Table 5.
Mtb RNAP core for in vitro reconstituted holoenzymes
To obtain the Mtb RNAP core complex (β, β’, His-tagged a, and
𝜔
), E. coli BL21(DE3)
cells were transformed with plasmids pET-Duet-rpoB-rpoC (encoding the β and β’ subunits) and
pAcYc-HisrpoA-rpoZ (encoding the N-terminal 10X His-tagged a and
𝜔
"subunits) and were grown
in LB media at 37oC to an OD600 of 0.6–0.8. Protein expression was then induced with 0.25 mM
IPTG and grown overnight at 16°C. Cells pellets resuspended in 20 mM Tris, pH 8.0, 5 mM
Imidazole, 500 mM NaCl, 5 mM β-ME with 1 protease inhibitor cocktail tablet (Roche,
05892791001) and lysed via sonication. Clarified cell lysate (via centrifugation at 5,000 rpm for
30 min at 8oC) was loaded on two 5 ml Ni2+ HisTrap FF crude affinity columns (Cytiva, 17528601)
set up in tandem using a 5–1000 mM imidazole gradient in 20 mM Tris, pH 8.0 and 500 mM NaCl.
Fractions of interest were dialyzed overnight (25 kDa MWCO tubing, Spectrum Labs, 132554) in
10 mM Tris, pH 8.0, 250 mM NaCl, 0.1 mM EDTA, 1 mM MgCl2, 10 μM ZnCl2, and 10 mM DTT,
concentrated, and further purified by size exclusion chromatography (HiPrep 16/60 Sephacryl S-
300 column, Cytiva, 17116701). Eluted complexes were dialyzed overnight in 10 mM Tris, pH 8.0,
200 mM NaCl, 0.1 mM EDTA, 1 mM MgCl2, 20 μM ZnCl2, 50% (v/v) glycerol, and 2 mM DTT,
defined here as storage buffer. Mtb RNAP core was concentrated to ~5 μM (100 kDa MWCO
Vivaspin 20 filters, Sartorius, VS2041), flash frozen in liquid nitrogen and stored at –80oC.
Concentration was determined with A280 with an extinction coefficient of 245,000 M-1 cm-1.
Co-expressed/purified Mtb σA RNAP holoenzyme
Co-expression/purification of the Mtb RNAP σA holoenzyme complex was obtained in the
following two ways: 1) use of a 10X N-terminal His-tag on a using pET-Duet-rpoB-rpoC, pAcYc-
HisrpoA-rpoZ, and pAC27-sigA plasmids and 2) use of a 10X N-terminal His-tag on σA using pET-
Duet-rpoB-rpoC, pAcYc-HissigA-rpoA, and pCDF-rpoZ plasmids (used only in 2 experiments in
the comparison of rrnA promoters section with no difference in activity). Complexes were
expressed and purified as described for Mtb RNAP core above. In some cases, an anion
exchange chromatography step (MonoQ 10/100 GL, Cytiva, 17516701) was added. Fractions of
interest were dialyzed overnight into 20 mM Tris, pH 8.0, 150 mM NaCl, 0.5 mM EDTA, 1 mM
MgCl2, 5% (v/v) glycerol and 2 mM β-ME and purified using 150–1000 mM NaCl gradient. The
final holoenzyme fractions were dialyzed into storage buffer, concentrated and frozen as
described for Mtb RNAP core, using an extinction coefficient of 280,425 M-1 cm-1.
Co-expressed/purified Mtb σB RNAP holoenzyme
Mtb RNAP σB holoenzyme complex was purified from a 10X N-terminal His-tag on a
construct (pET-Duet-rpoB-rpoC, pAcYc-HisrpoA-rpoZ , and pAC27-sigB plasmids) transformed in
E. coli NiCo21(DE3) cells (NEB, C2529H, (116)). Cells were grown at 25oC to an OD600 of 0.6,
induced with 0.5 mM IPTG and grown overnight at 16°C. Resuspension of pellets and protein
purification was carried out identically to that described for Mtb RNAP core (Ni2+ HisTrap FF crude
affinity followed by size exclusion chromatography), followed by anion exchange chromatography
as described for the Mtb σA RNAP holoenzyme. The holoenzyme was dialyzed into storage buffer,
concentrated and frozen as described for Mtb RNAP core, using an extinction coefficient of
264,955 M-1 cm-1.
Mtb σA, Mtb σB and Mtb σAΔ205
Mtb σA and Mtb σAΔ205 were purified from pET-SUMO plasmid vectors transformed in E.
coli BL21(DE3) and expressed as described for Mtb RNAP core above. Pellets were resuspended
in 50 mM sodium phosphate, pH 8.0, 5 mM imidazole, 300 mM NaCl, 5 mM β-ME with 1 protease
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inhibitor cocktail tablet and lysed via sonication. After centrifugation (5,000 rpm for 30 min at 8oC),
clarified lysate was incubated with Ni2+ NTA agarose beads (GoldBio, H350-50) for 30 min. The
column was washed with 20 mM imidazole and protein was eluted with 250 mM imidazole in 50
mM sodium phosphate, pH 8.0, 300 mM NaCl. His-SUMO tag was cleaved by Ulp1 during
overnight dialysis in 20 mM Tris, pH 8.0, 150 mM NaCl, 0.5 mM EDTA, 1 mM MgCl2, 5% (v/v)
glycerol and 2 mM β-ME in 6–8 kDa MWCO tubing (Spectrum Labs, 132655). The σ factors and
cleaved His-SUMO tag were then separated and further purified by anion exchange
chromatography (MonoQ 5/50 GL, Cytiva, 17516601) using a 150–1000 mM NaCl gradient.
Mtb σB was purified similarly, but with the following changes. E. coli NiCo21(DE3) cells
were used. Cell pellets were resuspended and sonicated in 50 mM potassium phosphate, pH 8.0,
20 mM imidazole, 500 mM ammonium chloride, 10% (v/v) glycerol, 0.1% triton X, 10 mM β-ME
with 1 protease inhibitor cocktail tablet. Clarified lysate was loaded onto a Ni2+ HisTrap FF crude
affinity column and protein was purified using a 5–1000 mM imidazole gradient in 20 mM
potassium phosphate, pH 8.0 and 250 mM ammonium chloride. The dialysis buffer for Ulp1
cleavage was 20 mM potassium phosphate, pH 8.0, 250 mM ammonium chloride and 1 mM β-
ME. The cleaved protein and the His-SUMO tag were then separated by an additional round of
Ni2+ HisTrap FF crude affinity chromatography, where σB was collected from the flow-through.
All sigmas were then dialyzed into storage buffer as described for RNAP core and
concentrated to ~40 μM (5 kDa MWCO Vivaspin filters, Sartorius, F27335) using extinction
coefficients of 35,410 M-1 cm-1 for Mtb σA, 29,910 M-1 cm-1 for Mtb σAΔ205, and 19,940 M-1 cm-1 for
Mtb σB.
Mtb CarD and RbpA
Mtb CarD and RbpA, in pET-SUMO plasmid vectors, were expressed, purified, and the
His-SUMO tag was cleaved in accordance with methods previously described (27, 36). In some
cases, after cleavage and separation of the SUMO tag, an additional anion exchange
chromatography step (MonoQ 10/100 GL, Cytiva, 17516701) was implemented using a 100–1000
mM NaCl gradient. Eluted fractions were then dialyzed overnight in 20 mM Tris, pH 8.0, 150 mM
NaCl, 1 β-ME, followed by concentration to ~200 μM determined using extinction coefficients of
16,900 M-1 cm-1 for Mtb CarD and 13,980 M-1 cm-1 for Mtb RbpA.
Plate-reader fluorescence measurements
For all experiments, data was collected using a CLARIOstar Plus Microplate reader (BMG
LabTech) in a 384 well, low volume, round-bottom, non-binding polystyrene assay plate (Corning,
4514) with the corresponding Voyager analysis software. To measure multi-round and single-
round transcription kinetics in real-time, we monitored the change in DFHBI fluorescence upon
binding to a transcribed, full-length RNA sequence containing the iSpinach D5 aptamer. DFHBI
fluorescence was measured with a monochromator excitation of 480 ± 15 nm, and the resulting
emission signal was monitored at 530 ± 20 nm. To measure dissociation of promoter-bound
complexes, Cy3 fluorescence was measured with a monochromator excitation of 535 ± 30 nm,
and the resulting emission signal was monitored at 585 ± 30 nm. All reactions were at 10 μl final
volume following initiation with 2.5 μl (single- and multi-round experiments) or 5 μl (dissociation
experiments) from automated reagent injectors (BMG LabTech). 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 37°C), 40 mM NaCl, 75 mM K-glutamate, 10 mM MgCl2, 5
μM ZnCl2, 20 μM EDTA, 5% (v/v) glycerol (defined as transcription buffer) with 1 mM DTT and
0.1 mg/ml BSA. All experiments were conducted at 37°C. For all experiments presented,
independent preparations of Mtb holoenzymes were used.
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Multi-round transcription experiments
Transcription reactions were performed with rNTPs (Thermo Scientific, R0481) at 1 mM,
3,5-difluoro-4-hydroxybenzylidene imidazolinone (DFHBI) dye (Sigma Aldrich, SML1627) at 20
μM, and RiboLock RNase inhibitors (Thermo Scientific, EO0381) at 0.4 U/μl. DFHBI concentration
was determined using an extinction coefficient at 420 nm of 31,611 M−1 cm−1. When applicable,
CarD and RbpA were added at concentrations of 1 μM and 2 μM, respectively. Transcription
reaction master mixes contained 75% of the final volume which included the RNAP holoenzyme
(co-expressed/purified or in vitro reconstituted), DFHBI, and RNase inhibitors, CarD/RbpA (when
applicable), and either DNA or rNTPs, depending on how the reaction was initiated. For
experiments comparing rrnA promoters and effects of DNA topology, master mixes containing
100 nM RNAP co-expressed/purified holoenzymes and 5 nM circular plasmid DNA templates
along with all other components were preincubated for 15 min at 37°C before initiation with rNTPs.
For σ titration experiments, 100 nM RNAP core was pre-incubated with various concentrations of
σ for 15 min at 37°C along with all other reaction components before initiation with 5 nM rrnAP3
circular plasmid DNA template. Data was typically acquired in 10–20 s intervals, not exceeding
40 min total. A minimum of 3 technical replicates of the negative control (leaving out DNA or rNTP)
were collected and measured concurrently with the experimental data. Using the average of this
negative control, the experimental data was corrected as previously described (37), bringing all
starting fluorescence values to zero and correcting for any time-dependent drift in fluorescence.
Between 4 and 7 independent experiments were collected for each condition with 3 technical
replicates each. Standard deviations were used as a statistical weight during the linear regression
analyses as previously described to obtain the steady-state rate (37).
Single-round transcription experiments
To promote single-round conditions by preventing dissociated/terminated RNAPs from
rebinding the promoter template, salmon-sperm DNA (Thermo Fisher Scientific, 15632011) was
used as a competitor. Competitor DNA was buffer exchanged in transcription buffer and
concentration was determined by A260. Promoter-bound complexes were pre-formed with 5 nM
rrnAP3 circular plasmid DNA and either 250 nM σA RNAP co-expressed/purified holoenzyme
supplemented with 2.5 μM σA or 125 nM σB RNAP co-expressed/purified holoenzyme
supplemented with 1.25 μM σB along with 20 μM DFHBI and 0.4 U/μl RNase inhibitors. These
concentrations were chosen to maintain the same protein:DNA ratio as used in the stopped-flow
assays (below). Following a 15 min incubation at 37°C, reactions were initiated with 25 μg/ml of
salmon-sperm DNA and 1 mM rNTPs. When competitor was first pre-incubated with RNAP
holoenzyme and rrnAP3 circular plasmid DNA containing the aptamer sequence, no change in
fluorescence upon initiating the reaction with NTPs was observed (Figure 4B). Data was acquired
in 6 s intervals, for 20 min total and underwent the same subtractions as described for the multi-
round experiments. The averages and standard deviations for 3 independent experiments are
presented.
Dissociation of promoter-bound complexes
Dissociation of RNAP-promoter bound complexes experiments were adapted from
previous methods described using stopped-flow rapid mixing (31) and those described below.
Briefly, in a final volume of 10 µl, 100 nM σA RNAP co-expressed/purified holoenzyme or 50 nM
σB RNAP co-expressed/purified holoenzyme supplemented with 500 nM σB and with 2 μM CarD/4
μM RbpA (when applicable) was incubated with 2 nM linear rrnAP3 promoter Cy3-labeled DNA
at 37◦C for 15 min. RNAP dissociation was measured by subsequent equal injection with 50 μg/ml
salmon-sperm DNA. All concentrations listed are those prior to equal volume dilutions due to
mixing. Data was acquired in 90 s intervals for a total of 20 hours. To avoid sample evaporation
of the sample over time, immediately following injection of competitor DNA, wells were covered
with an optical adhesive (Applied Biosystems, 4360954). Data was normalized to obtain the
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(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made
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fraction of the initial signal remaining at a given time with the formula: [(F – Fo)/(Fstart – Fo)] where
Fo is the buffer subtracted value for Cy3-labeled DNA alone, F is the buffer subtracted signal, and
Fstart is the buffer subtracted signal at the time of injection. The averages and standard deviations
for 3 independent experiments are presented, where the standard deviation was used as a
statistical weight in fitting the time-courses to a single exponential function for the σA holoenzyme
and a double exponential function for the σB holoenzyme.
Stopped-flow fluorescence measurements
For all experiments, data was collected at 37°C using a SX-20 stopped-flow
spectrophotometer (Applied Photophysics) with excitation provided by a 535 nm fixed-wavelength
LED light source with a 550 nm short pass cut-off filter (Applied Photophysics), while monitoring
emission using a 570 nm long pass cut-off filter (Newport Optics), as previously described (31).
Following equal volume mixing with a total shot volume of 100 μl, the final reaction conditions
corresponded to those of the transcription buffer with 1 mM DTT, and 0.1 mg/ml BSA. For all
experiments presented, independent preparations of Mtb holoenzymes were used, where
multiple technical replicates were combined to measure a ‘shot average’. The shot averages from
each holoenzyme preparation were weighted equally in determining the averages and errors
presented. Data was collected for 1000 or 2500 s with logarithmic sampling over 2500 points.
Open complex formation
For experiments comparing the basal kinetics of Mtb holoenzymes (Figure 3A), 50 nM σA
RNAP co-expressed/purified holoenzyme and 50 nM σB RNAP co-expressed/purified
holoenzyme supplemented with 500 nM σB were incubated at 37oC for 5 min prior to equal volume
mixing with 2 nM linear rrnAP3 promoter Cy3-labeled DNA. Experiments in the presence of 2 μM
CarD and 4 μM RbpA were conducted in the same way except that the σA RNAP co-
expressed/purified holoenzyme concentration was increased to 100 nM to saturate the equilibrium
fluorescence value (compare end point fluorescence value Figure 3A to Figure 5A). All
concentrations listed are prior to equal volume mixing. Data is plotted as fold-change over DNA
alone according to the formula: (F – Fo)/Fo, where Fo is the buffer subtracted value for Cy3-labeled
DNA alone and F is the buffer subtracted signal.
Promoter escape
100 nM σA RNAP co-expressed/purified holoenzyme or 50 nM σB RNAP co-
expressed/purified holoenzyme supplemented with 500 nM σB ± CarD/RbpA were pre-incubated
with linear Mtb rrnAP3 promoter Cy3-labeled DNA and were rapidly mixed with 50 μg/ml salmon-
sperm competitor DNA and 2 mM rNTPs (concentrations listed prior to equal volume mixing). As
both the dissociation and promoter escape assays are conducted under single-round conditions,
and as a result independent of RNAP holoenzyme concentration, these holoenzyme
concentrations were chosen since the open-complex equilibration signal is saturated, both in the
absence and presence of CarD/RbpA (Figure 5A,B).
Dissociation of promoter-bound complexes
Concentrations used for monitoring RNAP dissociation are identical to those described
above for dissociation measurements using the plate-reader assay. Here an equal volume of 50
μg/ml salmon-sperm competitor DNA was mixed with the co-expressed/purified holoenzyme, and
the linear rrnAP3 promoter Cy3-labeled DNA.
Acknowldgements
The authors would like to thank Dr. Jerome Prusa for discussions at the outset of this work, Abby
Tang for assistance with aptamer assay optimization, and Dr. Eric Tomko for critical reading of
the manuscript.
.CC-BY-NC-ND 4.0 International licenseavailable under a
(which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made
The copyright holder for this preprintthis version posted February 27, 2025. ; https://doi.org/10.1101/2025.02.24.639987doi: bioRxiv preprint
Authors contributions: Ana Ruiz Manzano: Conceptualization, Investigation, Methodology, and
Writing—original draft. Drake Jensen: Conceptualization, Investigation, Methodology, Validation,
and Writing. Eric Galburt: Conceptualization, Funding acquisition, Methodology, Supervision,
Validation, Visualization and Writing—original draft.
Funding
National Institutes of Health [R35GM144282 to E.A.G.] and Biochemistry and Molecular
Biophysics department [Seed Grant to A.R.M].
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