Structural basis of pausing during transcription initiation in mycobacterium tuberculosis

Domain: neuroscience
Published: 2026-01-30
Editorial outcome: major_revision

Abstract
In bacteria, RNA polymerase (RNAP) often pauses during the early stages of transcription initiation. The structural basis for these transient pauses remains unclear. Here, we present cryo-electron microscopy (cryo-EM) structures of the paused initiation complex (PIC) and initiation complex (IC) of Mycobacterium tuberculosis (Mtb), which include the RNAP core enzyme, the ECF σ factor σE, transcription factor CarD, promoter DNA, and nascent RNA. Our structures with pre-melted scaffolds reveal an intermediate at the 6–7 nt stage compatible with a paused-like intermediate, associated with steric hindrance between the emerging RNA and the σ3.2 region. This clash triggers a swivel of the RNAP structural module and scrunching of the transcription bubble. We also observe positional rearrangement of the σ4 domain, suggesting a poised pre-escape state. In addition, complementary reconstructions with fully matched DNA scaffolds (N-IC and N-PIC) support the physiological relevance of the captured intermediates. Together, our results support the existence of a mechanistic checkpoint during transcription initiation and suggest an RNA-induced model how RNAP conformational dynamics regulate early transcription.

Introduction
Transcription initiation is the first and critical step in gene expression across all kingdoms of life1–5. Bacterial RNA polymerase (RNAP) employs a σ factor to recognize gene promoter regions, unwind double-stranded DNA (dsDNA) to form a transcription bubble, facilitate the synthesis of initial short RNA transcripts, and ultimately either assists in promoter escape or results in abortive transcription5–7. Abortive transcription frequently causes pausing and release of short RNA products through stressed initiation complex (IC) intermediates during the early stages of transcription initiation8–10. Although numerous single-molecule fluorescence resonance energy transfer (sm-FRET) studies and biochemical experiments have been used to study the dynamic process of transcription initiation11–13, our understanding of initial transcription, especially pausing during transcription initiation, remains limited because of the transient nature of intermediates and molecular heterogeneity. In addition, the dynamic nature of the initial intermediate complexes poses a challenge for structural definition. Consequently, these pausing states from transcription initiation have not been captured in high-resolution structures.

Extra-cytoplasmic function (ECF) σ factors are the largest class of alternative σ factors related to primary σ factors (σA), comprising only three conserved modules (σ2 domain, σ4 domain, and the functional σ3.2 region), in contrast to σA, which contains σ1.1 region, σ1.2 region, σ2 domain, σ3 domain, σ3.2 region, and σ4 domain14–19. The σ2 and σ4 domains recognize the DNA promoter −10 element and −35 element, respectively20–22. The σ3.2 region plays several crucial roles, including linking σ2 to σ4, inserting into the RNAP active-center cleft to initiate de novo transcription, adopting a helical conformation to pre-organize the template single-stranded DNA (ssDNA), and occupying the path of nascent RNA to mimic an RNA molecule for interaction with the template ssDNA20–23. However, as RNA synthesis proceeds, the σ3.2 region must be displaced due to steric interactions with the 5’-end of nascent RNA24–26. This displacement imposes energy barriers that contribute to pausing and abortive initiation23,27–31, which are subsequently accommodated by scrunching: RNAP pulls the unwound single-stranded template DNA inward, accommodating the additional unwound DNA by creating kinks in the transcription bubble28,29,32. Scrunching allows RNAP to capture free energy from DNA stacking. This energy is subsequently stored and used to disrupt RNAP-promoter interactions, enabling promoter escape28,29. In summary, we selected the ECF σE for structural analysis because it has a simpler domain organization (comprising σ2, σ3.2, and σ4) than the primary σA. This minimal architecture made it experimentally more tractable for reconstitution and structural determination, enabling us to better capture initiation intermediates, including the paused complex.

The Mtb σE factor is a critical ECF σ factor that mediates responses to the cell-surface and other extra-cytoplasmic stresses5,14,33–39. Although σE shares high sequence homology with σH and is predicted by AlphaFold to adopt a highly similar fold, direct experimental structural information for Mtb σE factor is still limited. Recently Yu Zhang and Richard H. Ebright groups have reported the structures of Mtb transcription initiation complexes comprising the ECF σ factors σH and σL, respectively40,41. σH and σL adopt homologous binding modes to the RNAP core and promoter DNA, recapitulating the σ-dependent holoenzyme architecture observed in bacterial transcription initiation25,40,41. These structures also revealed that the σ3.2 linker of ECF σ factor is inserted into the RNA exit channel and active center cleft, playing an essential role in open promoter complex (RPo) formation, promoter escape and abortive production25,33,40,41. In addition to σE, CarD is a transcriptional activator found widely across bacterial species, including Mtb, but is absent in E. coli42,43. The structure of CarD consists of an N-terminal domain that folds into a Tudor-like structure, which is similar to the E. coli transcription repair coupling factor, and serves as the RNAP-interacting domain and a C-terminal helical domain42,43. In Mtb, CarD has been described as a regulator of the principal σ subunit, particularly in ribosomal gene transcription44, whereas its cooperation with ECF σ factors has not been reported. Notably, CarD has been reported to modulate transcription driven by an ECF σ factor in Myxococcus xanthus, suggesting that CarD may also influence σ factor–dependent transcription in Mtb, although this has not been experimentally characterized45. Thus, while the general mechanism by which ECF σ factors initiate transcription is increasingly well understood, the basis of transcriptional pausing during initiation remains poorly defined, in part due to limitations of traditional crystallographic approaches, and the potential involvement of CarD in this process has yet to be elucidated.

Here, we investigated the mechanism of pausing during transcription initiation by determining a series of cryo-electron microscopy (cryo-EM) structures of transcription complexes from Mtb, comprising the RNAP core enzyme, DNA/RNA scaffolds, ECF σ factor σE, and general transcription factor CarD. These complexes include (1) an initiation complex with 6-nt RNA (IC6), (2) an initiation complex with 7-nt RNA (IC7), and (3) a paused initiation complex with 7-nt RNA (PIC7). Our structures are consistent with RNA polymerase pausing during the transition between 6 and 7 nt transcripts, adopting a half-translocated conformation that swivels the RNAP, resulting in a 3-base pair unwinding and scrunching of the transcription bubble DNA. We also found that CarD binds to the template ssDNA in the IC to help maintain the transcription bubble, and interacts with the non-template ssDNA in the PIC to stabilize the bubble and potentially facilitate escape from the paused state. This difference in binding modes indicates that CarD plays an important role not only in the formation of the transcription bubble but also in the regulation of pausing during transcription initiation. In addition to these high-resolution structures obtained with pre-melted scaffolds, we also reconstructed initiation complexes using fully complementary DNA scaffolds to assess physiological relevance. These include (4) an initiation complex with 6-nt RNA (N-IC6), (5) an initiation complex with 7-nt RNA (N-IC7), and (6) a paused initiation complex with 7-nt RNA (N-PIC7). Although resolved at lower resolution and without atomic modeling, these reconstructions are consistent with the pre-melted scaffold structures and support the validity of our scaffold design strategy. By combining analyses of these structures with biochemical experiments, we support the presence of a pausing checkpoint during transcription initiation, and provide an RNA-induced model to elucidate the mechanism underlying paused transcription initiation.

Results
Preparation of the PIC using a nucleic acid scaffold with a 6- or 7-nt RNA
The ICs containing Mtb ECF σ factors have been successfully prepared using nucleic acid scaffolds for structural studies40. According to kinetic analyses of RNAP transcription initiation, RNAP exit kinetics from initiation complexes stalled specifically at the 7-nt RNA transcript stage demonstrate delayed transition to elongation11. Additional research has also shown that the pause is encountered by RNAP after the synthesis of a 6-nt RNA12. To test whether Mtb RNAP dwells at this stage, we performed in vitro transcription assays using fully complementary DNA-RNA scaffolds. RNAP showed accumulation of transcripts at 6–7 nt, consistent with increased dwell time during extension from 6 to 7 nt, and supporting the idea that the RNA lengths used for our structural studies correspond to relevant early-initiation intermediates (Supplementary Fig. 1H, I). These biochemical studies are therefore consistent with RNAP dwelling at the 6–7 nt stage, in agreement with the structural capture of PIC-like intermediates. In these assays, we use the fully matched scaffold to relate the 6–7 nt products directly to the cryo-EM architecture, whereas pre-melted scaffolds are interpreted as qualitative support for the presence of a 6–7 nt checkpoint.

Based on precedents from structural studies of mycobacterial RNAP and ECF σ factors (Supplementary Table 1), we adopted pre-melted nucleic acid scaffolds for IC6, IC7, and PIC7, as these designs are widely used to stabilize transcription initiation intermediates beyond RPo and allow visualization of the interactions between nascent RNA and the σ3.2 region. To investigate the potential formation of PIC and the mechanism of pausing, we prepared transcription initiation complexes that contained RNAP-core, σE factor, CarD, and nucleic acid scaffolds with a 6- or 7-nt RNA. We tested the binding of σE factor and CarD to the RNAP-core using native electrophoretic mobility shift assays (EMSA), and found that adding σE factor and CarD to the RNAP-core pre-incubated with DNA/RNA scaffold resulted in an upward shift of the band, indicating the formation of a potential complex (Supplementary Fig. 1G). After collecting and processing 9518 cryo-EM micrograph movies in the transcription initiation complex with a 7-nt RNA, two major classes were identified: 55% of the particles belonged to IC, and 45% to PIC (Supplementary Fig. 2). This distribution likely reflects the relative stability and population of these two conformations under the experimental conditions, representing alternative paused states within the initiation checkpoint. Additional transient intermediates may exist but were not captured due to their low population and dynamic nature; these could include partially scrunched states, transiently backtracked complexes, or short-lived RNA-σ3.2 interactions, consistent with the idea that cryo-EM preferentially samples the most stable states. At a matched resolution of 3.3 Å, the IC7 reconstruction utilized 75,465 particles while the PIC7 employed 65,908 particles. (Supplementary Fig. 2 and Supplementary Table 2). Only the major class was identified by collecting and processing 2786 cryo-EM micrograph movies in the transcription initiation complex with a 6-nt RNA which belonged to the IC. At a resolution of 3.5 Å, the IC6 reconstruction employed 65,079 particles following the same processing workflow (Supplementary Fig. 3 and Supplementary Table 2). All transcription complexes showed high-quality densities for RNAP, the σ2 domain of σE, and DNA/RNA scaffold, except for the σ4 domain of σE and CarD, whose densities were relatively weak but still sufficient to position the main chain (Supplementary Fig. 2, 3).

In addition to these high-resolution structures obtained with pre-melted scaffolds, we also performed reconstructions using fully complementary DNA scaffolds to assess physiological relevance. These reconstructions yielded N-IC6, N-IC7, and N-PIC7 (Supplementary Figs. 4, 5 and Supplementary Table 3). Although the maps were at lower resolution and lacked continuous density for parts of the DNA/RNA scaffold, they nevertheless showed overall conformational features consistent with the corresponding pre-melted scaffold structures (Supplementary Fig. 4C). Because of the limited resolution and incomplete nucleic acid density, atomic models could not be built for these reconstructions. These results support the rationale of using pre-melted scaffolds and are consistent with the view that the captured IC and PIC states represent physiologically relevant intermediates. We therefore focused subsequent structural analyses on the high-resolution IC6, IC7, and PIC7 reconstructions, while using N-IC6, N-IC7, and N-PIC7 as supportive validation datasets.

Overall structure of IC and PIC
In the IC structure, which includes IC6 and IC7 that are nearly identical except for RNA length differences and σ3.2 tip conformational changes, CarD and σE maintain equivalent interactions with RNAP as those observed in other ECF σ factor structures40,41. CarD forms a minor groove wedge to stabilize template ssDNA in the transcription bubble (Fig. 1A, B). The σ2 domain of σE engages the clamp module of RNAP β’ subunit via a broad interaction interface (Fig. 1B). In contrast, the σ4 domain of σE encapsulates the tip helix of the RNAP β flap domain (βFTH) within a tightly fitting structural pocket (Fig. 1B). The structure of the IC also revealed multiple interactions in promoter recognition and promoter unwinding by σE. The σ4 domain of σE anchors to the major groove of the upstream DNA and reads the sequence of the −35 element of the upstream dsDNA (Fig. 1B and Supplementary Fig. 6A). The σ2 domain of σE interacts with the junction and fork of the -10 element of the upstream ds/ssDNA to unwind the promoter DNA, similar to other ECF factors (Fig. 1B and Supplementary Fig. 6A)25,40,41. Moreover, the σ3.2 region of σE inserts into the active center cleft to guide the template ssDNA into the RNAP active site (Fig. 1B). The DNA/RNA hybrid base pairs remain in a post-translocation state within the active site, with a root-mean-square deviation (RMSD) of 1.819 Å for all Ca atoms compared to the post-translocation state in Mtb (PDB: 5ZX2) (Fig. 1C, D and Supplementary Fig. 7A, B).

In the PIC structure, while the interactions of the σ2 and σ4 domains of σE with RNAP resemble those in the IC, multiple conformational changes differing from those observed in the IC are evident (Fig. 1A, E). Unlike the structure of IC, the PIC exhibited a half-translocated DNA/RNA hybrid, with an RMSD of 0.966 Å for all Ca atoms compared to the half-translocation state in Mtb (PDB: 8E8M) (Fig. 1F, G and Supplementary Fig. 7C, D). The swivel module (defined in Supplementary Table 4) rotated relative to the rest of the RNAP, resulting in rotation of the promoter upstream dsDNA (Fig. 1B, E). The upstream ds/ssDNA junction also undergoes unwinding of three bases, which may be caused by the combined effects of RNAP swiveling and the raising of the upstream dsDNA (Fig. 1B, E and Supplementary Fig. 6b). Moreover, the σ4 domain of σE, which anchors the major groove of the upstream dsDNA, shows a significant conformational change in PIC compared to IC (Fig. 1B, E). Surprisingly, CarD also formed a minor groove wedge but stabilized the non-template ssDNA in the transcription bubble (Fig. 1B, E).

The conflict between the σ3.2 and nascent RNA induces pausing of RNAP
The σ3.2 region of σE inserts into the active center cleft and occupies the RNA exit channel, which means it must be displaced as RNA elongates24–26. It has been reported that RNAP adopts a pausing state when the nascent RNA extends to 6 to 7 nucleotides, with the first evidence demonstrated in 201611–13. However, conformational changes in RNAP during pausing transcription initiation have not been directly observed. Additionally, RNAP swiveling, as a conserved feature, has been reported in previous studies on Mtb NusG-dependent pausing and the E. coli pause hairpin (PH) stabilized pausing elongation complex46–48. Despite these findings, it remains unknown whether RNAP swiveling occurs during pausing transcription initiation, what drives the change in the RNAP active site, and how the σ3.2 region of σE is displaced. Structural evidence from PIC7 provides key insights into these questions. The structure of PIC7 revealed that Mtb RNAP undergoes swiveling during pausing transcription initiation (Fig. 2A). Swiveling associated with pausing transcription initiation is similar to Mtb NusG-dependent pausing, which is involved in transcription elongation and termination, suggesting that swiveling may be a conserved characteristic in all pausing processes of transcription46–48. The PIC was swiveled and half-translocated, whereas the IC was unswiveled and post-translocated (Fig. 2A and Supplementary Fig. 7A–D). In the half-translocation state, the RNA moves forward, but the template DNA is not completely translocated. The swiveling of RNAP also distorts the bridge helix (BH), which is consistent with the known ability of Mtb NusG to facilitate pausing through interaction with the conserved TTNTTT motif46–48. The C-terminal portion of BH was attached to the swivel module, whereas the N-terminal portion of BH was attached to the unswivel module of the RNAP (Fig. 2B). Therefore, the middle of BH is kinked and the C-terminal portion of BH is shifted toward the template DNA, resulting in steric clash with the +1 position of template DNA in the nucleotide addition cycle (Fig. 2B).

Central to pausing maintenance is the σ3.2 region’s obstruction of RNA extension. The nascent RNA encountered a roadblock formed by the σ3.2 region of σE, which drives RNAP swiveling and maintains the pausing state. Although the swiveling associated with pausing is a conserved characteristic in all pausing processes of transcription, the reasons for driving swiveling in each pausing process are distinct. The E. coli his RNA hairpin stabilizes pausing through its interaction with the RNA exit channel, while Mtb NusG facilitates pro-pausing through the interaction of its N-terminal NGN domain with a conserved TTNTTT motif46–48. However, in Mtb σE-dependent pausing, the σ3.2 region of σE inserts into the RNA exit channel, forming a steric roadblock that clashes with the extending RNA. This steric interference promotes RNAP swiveling and contributes to transcriptional pausing, as supported by our in vitro transcription assays (Fig. 2B–E, Supplementary Fig. 1H–K), which reveal that pausing occurs during the transition from 6-nt to 7-nt RNA synthesis. This behavior is consistent with previous observations in the E. coli σ⁷⁰-RNAP system11.

Conformational restructuring of σE further elucidates pausing regulation. In the structures of IC6 and IC7, the σ2 and σ4 domains of σE are located on the surface formed by RNAP, while the σ3.2 region of σE, which likely coincides with the movement of the 5’-end of RNA and causes steric clash between the nascent RNA and the tip of the σ3.2 region, is inserted into the RNA exit channel (Fig. 2C–E). In the structure of PIC7, the folding of σ2 of σE is virtually identical to that in IC6 and IC7, while the σ4 domain of σE, which binds to the major groove of the upstream dsDNA, exhibits a remarkable conformational change within its relative position (Supplementary Fig. 8A–C). The σ4 domain in PIC7 rotates ~55° relative to its position in IC6, moving from the RNA exit channel region toward the upstream dsDNA. (Fig. 1B, E and Supplementary Fig. 8A). In IC and other ECF σ complexes, the σ4 central region is positioned 10 Å from the RNA exit channel25,40,41 (Fig. 1B, E). Upon transition to the PIC, σ4 central region undergoes rotation to a position 20 Å away from the channel, driving dissociation of the σ3.2 region from the β C-terminal helix (Fig. 1B, E). This previously unobserved position of the σ4 domain of σE not only causes variation in the σE and RNAP interactions, but also leads to significant differences in σE and DNA interactions (Supplementary Fig. 6A, B). It is worth noting that despite low sequence conservation in the σ3.2 region across ECF σ factors, we observe striking structural conservation of their flexible linker conformations (Supplementary Fig. 8A, D). This suggests a convergent functional mechanism for transcriptional pausing regulation, potentially mediated through σ3.2-dependent steric clashes during RNA extension.

The upstream DNA scrunching, unwinding, and raising in PIC
Before the promoter escapes, RNAP must provide sufficient time for DNA scrunching to accumulate the necessary free energy28–30. The highly stable scrunched state and extensive pausing during transcription initiation have been observed using sm-FRET11,12,25; however, the structure of pausing during transcription initiation, involving a scrunched, unwound, and raised state, has not been obtained. Our structural determination of the PIC reveals a 3-bp unwound region at the upstream ds/ssDNA junction, contrasting with the fully duplexed state observed in the IC (Fig. 3A, B). The driving force of unwinding may come from the compression of nascent RNA against the σ3.2 region, leading to the lifting of the σ4 domain and the upstream dsDNA, which in turn further promotes DNA unwinding (Fig. 3A–C).

The unwinding in PIC causes the scrunching of DNA in the transcription bubble, in which the upstream dsDNA rotates approximately 35° around the ds/ssDNA junction relative to its position in the IC structure, resulting in the −35 element of the upstream dsDNA being positioned at an increased distance of approximately 20 Å from the RNA exit channel (Fig. 1E). Although the pre-melted DNA scaffold used in our study facilitates the stabilization of transcription intermediates suitable for structural determination, it may result in a lower degree of scrunching compared with that occurring during the actual transcription reaction. The true scrunched states are likely to be more compact and dynamic, but their intrinsic instability currently prevents high-resolution structural characterization. Nevertheless, the paused conformation we observe here is consistent with that seen using fully matched DNA templates, supporting that the captured structure faithfully reflects the pausing state of the transcription complex (Supplementary Figs. 4, 5). Although the DNA duplex undergoes unwinding and rotational adjustments in PIC, extensive interactions persist between the PIC-associated DNA/RNA scaffold and the RNAP-core. However, the spatial arrangement of these interaction sites differs significantly compared to those observed in the IC (Supplementary Fig. 6A, B). In PIC assembly, the σ4 domain of σE and the major groove of dsDNA of the −35 element are adjacently located, unlike the tight binding observed in the IC, but they still separate and form a clamp of a narrow cleft with a width of approximately 10 Å (Supplementary Fig. 8B, C). The σ4 domain helix spanning from K221 to L240 engages the −35 element through weak van der Waals interactions, with the closest contact of approximately 5 Å localized between the helical surface and the T-26 to T-28 bases (Fig. 3C, D and Supplementary Fig. 8B, C). This minimal interface suggests a transient association mechanism at the transcriptional checkpoint, i.e., the stage where RNAP either pauses or escapes into productive elongation, distinct from stable binding. The reduced σE-DNA binding and transcriptional activity observed with R226A, R228A, and R231A substitutions could involve impaired recognition of the −35 element by the σ4 domain, along with modified structural rearrangements associated with σ4 release from the RNAP-core (Fig. 3E, F and Supplementary Fig. 8E). Thus, the IC and PIC represent two alternative conformations of RNAP at the 7-nt RNA stage, corresponding to distinct states within a transcriptional checkpoint.

CarD stabilizes distinct ssDNA in both IC and PIC
CarD interacts directly with the RNAP β subunit β1-lobe and flanks one side of the upstream ds/ssDNA junction, facing the β’ coiled-coil and the σ2 domain42. In both IC and PIC structures, CarD exhibits a significant reconfiguration in DNA-binding interactions, despite maintaining virtually identical folding with an all-Cα RMSD of 1.826 Å (Fig. 4A–E). In the structure of IC, the C-terminal helical domain of CarD directly interacts with the template DNA through two α-helices, using the hydrophobic planar side chain of W85 as a bulky wedge inserted into the minor groove at the junction of the template ssDNA (Fig. 4A, B), which is identical to that in previously reported CarD-RPo complexes42. The two α-helices were oriented approximately perpendicular to the dsDNA axis, establishing a relatively modest interaction interface of 380 Å2 between CarD and the template ssDNA (Fig. 4A, B and Supplementary Fig. 9A–C). In the PIC structure, the C-terminal helical domain of CarD directly engages the non-template ssDNA via two α-helices (Fig. 4C, D). This interaction is stabilized by potential hydrogen bonds formed between the side chains of Y89 and R126 with the phosphate groups of the DNA backbone, creating an interface between CarD and the ssDNA that resembles the template-strand interaction observed in the IC (Fig. 4C, D). CarD flanks one side of the non-template ssDNA, while the σ2 domain flanks the opposite side, together forming a tighter interaction with the non-template ssDNA to further stabilize the scrunched transcription bubble (Fig. 4C, D and Supplementary Fig. 9D–F). This stabilization likely prevents premature collapse of the bubble, while allowing the conformational flexibility required for RNAP to progress through the paused intermediate.

Surprisingly, structural comparison reveals that the PIC-specific ordering of the σ2 N-terminal helix correlates with CarD positioning near σ2, suggesting that CarD stabilizes this region via direct spatial interactions (Fig. 4F). Consistent with these structural observations, our in vitro transcription assays showed that CarD enhances σᴱ-dependent transcription and alters the distribution of RNA products at the 6–7 nt stage (Supplementary Fig. 1H–K), suggesting that CarD may play a dual role by stabilizing RNAP-σ-DNA complexes and potentially facilitating the transition from the paused to the elongating state during transcription initiation. Moreover, a time-resolved fluorescence assay using a Cy3-labeled promoter bubble was performed to assess the effect of CarD on open complex formation. In the presence of CarD, the fluorescence signal reached a higher plateau compared with the RNAP-σE-only complex, indicating that CarD stabilizes the σE-RPo complex and enhances its steady-state accumulation (Fig. 4G). These results are consistent with previous findings showing that CarD promotes the stabilization of transcription initiation complexes49,50. W85A, Y89A, and R126A mutations disrupted critical interactions, leading to a marked reduction in transcriptional activity (Fig. 4H). The structural studies of Mtb CarD-IC and CarD-PIC complexes presented here suggest that CarD plays an important role in forming the transcription bubble in the IC and in stabilizing the scrunched transcription bubble in the PIC.

Methods
Protein production and purification
The Mtb RNAP core enzyme was prepared by co-expression of genes for the Mtb RNAP α subunit, RNAP ω subunit, RNAP β′ subunit, and C-terminally deca-histidine-tagged RNAP β subunit in E. coli [plasmids pRSFDuet-rpoA-rpoZ and pETDuet-rpoB-rpoC]. The genomic material was purchased from BeNa Culture Collection. The Mtb RNAP core enzyme was expressed in E. coli BL21 (DE3) (Supplementary Table 5 and 6). The cells were harvested, resuspended in lysis buffer (20 mM Tris-HCl, pH 8.0, 150 mM NaCl, and 4 mM MgCl2), and homogenized with an ultra-high-pressure cell disrupter at 4°C. Cell debris was removed by centrifugation at 30,000 xg for 35 min. The Mtb RNAP core enzyme was first purified by Ni-NTA (Novagen, USA) affinity chromatography and then further purified by using a Hitrap Q ion-exchange column (GE Healthcare, USA) with buffer A (20 mM Tris-HCl, pH 8.0, 4 mM MgCl2, 2 mM DTT) and buffer B (20 mM Tris-HCl, pH 8.0, 1 M NaCl, 4 mM MgCl2, 2 mM DTT). Next, the sample was loaded onto a Superose 6 Increase 10/300 column (GE Healthcare, USA) with DEPC-treated buffer (20 mM HEPES, pH 7.0, 150 mM NaCl, 4 mM MgCl2, and 2 mM DTT). The purified Mtb RNAP core enzyme was concentrated to 10 mg/mL and stored at −80°C (Supplementary Fig. 1A-C).

The Mtb σE or its derivative gene was cloned into the pETDuet-1 vector with an N-terminal 6× His-tag. The genomic material was purchased from BeNa Culture Collection, and any mutations were introduced using primers listed in Supplementary Table 6. The protein was expressed in the E. coli strain BL21 (DE3) (Supplementary Table 5). Cells were grown at 37°C and induced at an apparent OD600 of 0.6–0.8 by addition of isopropyl-β-D-thiogalactoside (IPTG) to a final concentration of 0.2 mM, and then further grown at 16°C for 18 h. After harvesting by centrifugation, the cells were resuspended in lysis buffer (20 mM Tris-HCl, pH 8.0, 150 mM NaCl, and 4 mM MgCl2) and homogenized with an ultra-high-pressure cell disrupter at 4°C. The lysate was centrifuged at 30,000 xg for 35 min to remove cell debris. The fusion protein was first purified by Ni-NTA (Novagen, USA) affinity chromatography and then further purified through passage a Hitrap Q ion-exchange column (GE Healthcare, USA) with buffer A (20 mM Tris-HCl, pH 8.0, 4 mM MgCl2, 2 mM DTT) and buffer B (20 mM Tris-HCl, pH 8.0, 1 M NaCl, 4 mM MgCl2, 2 mM DTT). Next the sample was purified by application to a Superdex 75 10/300 Increase column (GE Healthcare, USA) with DEPC-treated buffer (20 mM HEPES, pH 7.0, 150 mM NaCl, 4 mM MgCl2, 2 mM DTT). Purified Mtb σE was concentrated to 10 mg/mL and stored at −80°C (Supplementary Fig. 1A-C).

Mtb CarD or its derivative gene was cloned into the pRSFDuet-1 vector. The genomic material was purchased from BeNa Culture Collection, and any mutations were introduced using primers listed in Supplementary Table 6. CarD with an N-terminal 6× His-tag was expressed in E. coli strain BL21 (DE3) (Supplementary Table 5). Cells were grown at 37°C and induced with 0.2 mM isopropyl-β-D-thiogalactoside at OD 600 nm 0.6–0.8. After induction at 16°C for 18 h, the cells were harvested and resuspended in lysis buffer (20 mM Tris-HCl, pH 8.0, 150 mM NaCl, and 4 mM MgCl2) and homogenized with an ultra-high-pressure cell disrupter at 4°C. After centrifugation at 30,000 xg for 30 min, recombinant protein was purified by Ni-NTA (Novagen, USA) affinity chromatography, and then was further purified by using a Hitrap Q ion-exchange column (GE Healthcare, USA) and a Superdex 75 10/300 Increase column (GE Healthcare, USA) with DEPC-treated buffer (20 mM HEPES, pH 7.0, 150 mM NaCl, 4 mM MgCl2, 2 mM DTT). Purified Mtb CarD was concentrated to 10 mg/mL and stored at −80°C (Supplementary Fig. 1A-C).

Assembly of the transcription (pausing) initiation complexes
Nucleic acid scaffolds for Cryo-EM complex assembly were prepared from synthetic oligos (Jiangsu GenScript Biotech Co., Ltd, Supplementary Fig. 1D, H and Supplementary Table 7) in DEPC-treated buffer (20 mM HEPES, pH 7.0, 150 mM NaCl, 4 mM MgCl2). Two strategies were used depending on scaffold type: (i) Pre-melted scaffold, Template DNA, non-template DNA, and RNA (The 5′ end of the RNA carries a phosphate group) were mixed at a 1:1:1 molar ratio and annealed by heating at 95°C for 5 min followed by stepwise cooling to 4°C. The resulting pre-melted scaffold was incubated with Mtb RNAP-core at a 1:1.2 molar ratio at 25°C for 10 min to form a stable RNAP-DNA/RNA complex. The mixture was then incubated with σE and CarD at a 1:3:3 molar ratio to assemble the σE_CarD_RNAP complex. The complexes were purified on a Superose 6 Increase 10/300 column (GE Healthcare, USA) equilibrated in the same buffer. Fractions containing Mtb RNAP-core, σE, and CarD were collected, concentrated to 6 mg/mL, and stored on ice until application to EM grids (Supplementary Fig. 1A-C; scaffold design adapted from ref. 34., with sequences shown in Supplementary Fig. 1D). (ii) Fully matched scaffold, Template DNA and RNA (The 5′ end of the RNA carries a phosphate group) were first annealed at a 1:1 molar ratio. The resulting tDNA-RNA duplex was then incubated with Mtb RNAP-core at a 1:1.2 molar ratio at 25°C for 10 min. Subsequently, non-template DNA was added at a 2-fold molar excess relative to the tDNA-RNA duplex and incubated at 25°C for 30 min, resulting in a stable RNAP-DNA/RNA complex. The mixture was then incubated with σE and CarD at a 1:3:3 molar ratio to assemble the σE_CarD_RNAP complex. The resulting complexes were loaded onto a Superose 6 Increase 10/300 column (GE Healthcare, USA) equilibrated in the same buffer. Fractions containing Mtb RNAP-core, σE, and CarD were collected, identified by SDS-PAGE (Supplementary Fig. 1E, F; gel: Cat No. Gel442012; Biotend, Shanghai, China), concentrated to 6 mg/mL, and stored on ice until application to EM grids (scaffold sequences are shown in Supplementary Fig. 1H and Supplementary Table 7).

Native electrophoretic mobility shift assays
The binding reaction buffer contained 20 mM HEPES, pH 7.0, 150 mM NaCl, 4 mM MgCl2. A total of 18 μg of Mtb RNAP core enzyme was combined with a 1.5 μg nucleic acid scaffold, and σE and CarD were mixed in a 1:2:2 molar ratio. Binding reactions were incubated for 25 min at 30°C. Samples were run on a 6% polyacrylamide native gel (37.5:1 acrylamide:bis-acrylamide) running in 1× TBE buffer at 100 V for 1.5 h in 4°C. The gel was stained with ethidium bromide.
The DNA binding affinities of σE was also quantified through EMSA using the same experimental procedure described above, with two modifications: (1) RNAP-core was omitted from the reaction system, and (2) the molar ratio of DNA to proteins σE was adjusted to 1:5.

Time-resolved fluorescence assay for RPo complex stabilization
The Cy3-labeled template and BHQ-labeled non-template (Supplementary Table 7), with fluorescent labeling performed as previously described49, were incubated with RNAP core enzyme at 37°C for 10 min at a 1:2 molar ratio in DEPC-treated buffer containing 20 mM HEPES (pH 7.0), 150 mM NaCl, 4 mM MgCl₂, and 2 mM DTT. σE and/or CarD were then added at a molar ratio of 1:5:5 (RNAP-core:σE:CarD) to assemble the σE_RNAP_DNA and σE_CarD_RNAP_DNA complexes. Fluorescence intensity was monitored using a Tecan Spark 20M multimode microplate reader (Tecan Group Ltd., Switzerland) at 25°C, with excitation at 510 nm and emission at 570 nm. Data points were collected every 3 s for 900 s. The fluorescence value of DNA-only control (FDNA) was subtracted as background, and the fluorescence fold-change was calculated as (Ft - FDNA)/FDNA. DNA-only, σE–RNAP–DNA, and σE–CarD–RNAP–DNA samples were analyzed in parallel. Fluorescence traces were background-corrected, normalized to the initial signal (F0), and fitted to a single-exponential association model, Ft = F0 + A(1 - e-kt), using GraphPad Prism 10 to obtain the apparent rate constant (k) and amplitude (A). All measurements were performed in triplicate, and data are reported as mean ± SEM from independent experiments.

Fluorescence-detected in vitro transcription assay
pCp-AF647-fluorescence-detected. the σE-dependent RNAP de novo transcription activity was measured by coupling pCp-AF647-labeled nucleotides to nascent RNA products75. Briefly, template DNA was used at a final concentration of 50 nM and incubated with Mtb RNAP core enzyme at a final concentration of 200 nM and σE at a final concentration of 400 nM in DEPC-treated buffer containing 20 mM HEPES (pH 7.0), 150 mM NaCl, 4 mM MgCl₂, and 2 mM DTT at 37°C for 15 min to assemble the holo-RNAP complex. CarD was then added at a final concentration of 1 µM, or omitted, and incubated for 5 min to assess its effect on transcriptional pausing. Single-round transcription was ensured by addition of heparin at a final concentration of 50 µg/mL. For reactions using the fully matched DNA template, RNA extension was allowed to proceed for 30 min at 37°C. For reactions using the pre-melted DNA template, aliquots were collected at 0, 15, 30, and 60 min for analysis. RNA products were extracted with Trizol reagent (VeZol Reagent: Vazyme Biotech Co., Ltd) and precipitated with isopropanol in the presence of GlycoBlue at a final volume of 1 µl at −20°C overnight. Precipitated RNA was washed twice with pre-cooled 70% ethanol, air-dried at room temperature, and resuspended in 2.5 µl RNase-free water. The RNA was then ligated with pCp-AF647 using T4 RNA ligase 1. Reactions were stopped by addition of 2× TBE-Urea loading buffer, and RNA-pCp-AF647 products were resolved on freshly prepared 20% urea-polyacrylamide gels containing 8 M urea. Fluorescence signals were detected using a GelView 6000 Pro II scanner. All experiments were performed in triplicate.

Mango-fluorescence-detected. transcription activity was measured by utilizing the significantly enhanced fluorescence of TO1-3PEG-Biotin when the Mango riboswitch was engaged, as previously reported76. Briefly, double-stranded nucleic acid scaffolds for fluorescence-detected in vitro transcription assays were prepared by PCR using pUC57-PsigB-Mango-tR2 as a template (Supplementary Table 7). The transcription activity of σE mutants or CarD mutants was determined in a 20 µL reaction mixture (20 mM HEPES, pH 7.0, 150 mM NaCl, 4 mM MgCl2, 2 mM DTT) containing the Mtb RNAP core enzyme (final concentration 80 nM), promoter DNA or its mutants (final concentration 100 nM), σE or its mutants (1 µM), and CarD or its mutants (1 µM) at 37°C for 10 min. The reaction was initiated by adding 2 µL NTP mixture (ATP, UTP, CTP, and GTP; final concentration 0.1 mM of each) and 2 µL TOl-3PEG-Biotin (final concentration 0.5 µM), and RNA synthesis was allowed to proceed for 30 min at 37°C. Fluorescence signals were measured using a Tecan Spark 20M multimode microplate reader (Tecan Group Ltd., Switzerland) with an excitation wavelength of 510 nm and emission spectra recorded 550 nm. All experiments were performed in triplicate and analyzed using GraphPad Prism 10.

Cryo-EM sample preparation and data collection
In total, 3.5 µL of protein solution at 6 mg/mL (added with 0.025% DDM, from Merck, supplied by Shanghai YuanXiang Medical Equipment Ltd, China) was applied on a glow-discharged R0.6/1.0 200-mesh Quantifoil grid (Quantifoil, MicroTools GmbH, Germany). The grid was then blotted for 4.5 s with a blot force of 0 at 8°C and 100% humidity, and plunge-frozen in liquid ethane using Vitrobot (Thermo Fisher Scientific, USA). The grids were loaded into a 300 keV Titan Krios electron microscope (Thermo Fisher Scientific, USA) equipped with a K3 direct electron detector (Gatan, USA). All images were automatically recorded using a SerialEM with a nominal magnification of 81000× and a calibrated super-resolution pixel size of 0.89 Å/pixel. The exposure time was set to 2.4 s with a total accumulated dose of 55 electrons per Å2. The images were collected with a defocus range from −1.0 to −1.8 µm. The statistics for the data collection and refinement are shown in Supplementary Table 2 and 3.

Cryo-EM image processing
For all samples, the raw movies were motion-corrected and dose-weighted using the MotionCorr2 v1.4.577 software, and their contrast transfer functions were estimated using ctffind4 v4.1.378. Particles were automatically picked using a blob picker and extracted with a box size of 320 pixels using cryoSPARC v4.5.179. The following 2D, 3D classifications and refinements were all performed in CryoSPARC. The particles were selected after two rounds of 2D classification based on complex integrity. This particle set was used to perform Ab-Initio reconstruction in eight classes, which were then used as 3D volume templates for heterogeneous refinement. Particle classes containing the complete σE-CarD-RNAP-DNA/RNA complex were retained for further refinement, while classes lacking one or more components were excluded. As a result, 75,465 particles converged into one Mtb IC7 complex class and 65,908 particles converged into one Mtb PIC7 complex class. Next, these particles were used for Non-uniform refinement to obtain one class of particles with final resolution of 3.3 Å. The same methods used for IC6 with a 6-nt RNA are described in Supplementary Fig. 2, 3. Similarly, the N-IC6, N-IC7, and N-PIC7 datasets were processed following the same workflow. After 2D and 3D classification, 25,048 particles for N-IC6, 15,872 particles for N-IC7, and 31,912 particles for N-PIC7 were retained for Non-uniform refinement, resulting in final reconstructions at 3.6 Å, 4.0 Å, and 3.9 Å resolution, respectively (Supplementary Fig. 4, 5).

Model building and refinement
To determine the structure of the Mtb IC complex and Mtb PIC complex, the structures of the Mtb RNAP core enzyme and σH complex (PDB: 5ZX2) and CarD (PDB: 6EDT) were individually placed and rigid-body fitted into the cryo-EM map using UCSF Chimera80. The DNA/RNA scaffold was manually built in Coot81 guided by the cryo-EM density and prior scaffold designs (PDB: 6JBQ)32. RNA positioning, especially near the σ3.2 region, was adjusted to match the observed density. The complete model was refined in real space using Phenix v1.19.282, with secondary structure and geometry restraints applied. Additional weight optimization was used to balance the fit between protein and nucleic acid components. Model validation was performed using MolProbity for geometry assessment and FSC between model and map for map-to-model fit. The DNA and RNA densities in the active site region were well-resolved, confirming accurate positioning and correct interactions with protein components. Final refinement statistics are provided in Supplementary Table 2.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This study reports the three cryo-EM structures of sigma E-RNAP transcription initiation intermediates. There are new\ninsights in the structure PIC7 (paused initiation complex with 7-nt RNA): (1) a half-translocated state of the DNA-RNA hybrid;\n(2) a swivel conformation of the RNAP resulting in shifted location of upstream dsDNA and the CarD. The author proposed\nthat the steric hindrance between sigma 3.2 and the nascent RNA is likely the causing factor inducing pausing at the\ntranscription initiation stage. Overall, this study presents an RNAP conformation that has not been observed before. The flaw\nin the method compromises the reliability of the conclusion. I have following concerns.\nMajor concerns:\n1. The major concern is the DNA-RNA scaffolds, which have been used to reconstitute the RPitc6 and RPitc7 complexes.\nThe authors did not describe in detail the logic for designing the DNA-RNA scaffolds. The best way to capture the RPitc6\nand RPitc7 is by adding subset NTPs to RPo (prepared by incubating RNAP with a fully complementary promoter DNA) and\nallowing RNA synthesis to 6 or 7 nt. If the method is not feasible, you can reconstitute the complexes with a DNA-RNA\nscaffold containing an RNA of 6 or 7-nucleotide and a double-stranded DNA, but you should use the complementary\nsequences for the non-template and template DNA and use an RNA with sequence precisely matching the nascent RNA\nproduced in vivo or in vitro transcription assay. The length of nascent RNA and the size of the transcription bubble are\nclosely related. For example, if the RPo has a transcription bubble of 12 bp, then an RPitc6 in post-translocation state should\nhave a transcription bubble of 17 bp, and an RPitc7 in half-translocated state should also have a transcription bubble of 17\n(RNA has translocated but DNA has not). So if the author would like to use the reconstituted complexes for structure\ndetermination, it is suggested first to identify the transcription start site of a promoter and then to determine an RPo structure\nwith the promoter as an unscrunched bubble control. For RPitc6 and RPitc7, the reviewer suggests designing scaffolds with\nthe correct RNA sequence to define the natural bubble size at the respective states. The current scaffolds have non-\ncomplementary sequences in the transcription bubble and incorrect bubble size for RPitc6 and 7, and therefore, they are not\nthe physiological-relevant transcription intermediate complexes.\n2.The transcription initiation complex with a 7-nt RNA yield two major conformation (IC and PIC). The authors should give\nsome explanation for why these two distinct conformations arise. Do these conformations represent sequential\nintermediates, or are they alternative pathways during transcription?\n3.Figure 3a,b-The authors should include zoomed-in views to highlight the contrasting regions of ‘unwinding’ and ‘non-\nunwound’ template DNA. The current version does not show this with enough clarity. The DNA should be labeled relative to\nthe TSS.\n4. The manuscript lacks a clear explanation of how the authors selected the particle classes of interest during cryo-EM data\nprocessing. It is recommended to clarify whether selection was based on resolution, particle percentages, or other criteria.\nPlease provide this information either in the Extended Data Figures or in the Methods section.\nMinor issues:\n1. Extended Data Fig. 2: The label “Mtb IC6” appears to be incorrect and should be revised to “Mtb IC7”.\n2. More detail is needed regarding the model building procedure, particularly for how the DNA and protein components were\nmodeled and validated.\n3. Line 474: “a 6 polyacrylamide native gel” should be “a 6% polyacrylamide native gel”.\n4. Figure 2d: The DNA template nucleotides in the active site appear to be missing.", "reviewer_2": "The work of Zheng et al provides structural snapshots of important intermediates of the transcription path from initiation to\nelongation, centred around the pause site that has been observed to occur after the synthesis of around 6-7 nt of RNA.\nDespite the available information biochemical and biophysical information, there are no high-resolution structures of the\npaused state. Zheng et al used cryoEM to provide such information for the Mtb system and reveal several interesting\nconformational changes, the most exciting being the presence of a half-translocated state with RNAP swiveling in the\npaused complex (which to my knowledge has not been observed before in initiation), a motion that has been associated with\ndifferent types of pausing in transcription elongation. This observation draw nice parallels between initiation and elongation\nand offers an additional element for the regulation of transcription initiation in bacteria, which may have generality beyond\nthe Mtb system.\nThe paper is overall well written, although there are points that needs attention, and aspects that needs addressing before\npublication.\nMajor points\n1. The authors need to be much more explicit about the fact that the complexes prepared are synthetic, i.e., not prepared\nduring actual synthesis, but assembled using synthetic nucleic acid scaffolds (Ext Fig 4) – as a result, it appears that the\ncomplexes have much less scrunching that actual complexes with the same RNA length would have had, and do not reflect\nexactly the final structures of complexes formed via actual RNA synthesis. For example, for the template provided, an IC7\nwould expect to melt the region from +1 to +7; however, currently the RNA is hybridized using the existing bubble. The\nauthors need to discuss how their main and secondary findings are affected by the assembly method, and why this\nassembly route was pursued.\n2. L62. Do we know that the picture for the ECF σE is similar as for σA? Why was σΑ not selected to proceed with structural\nanalysis?\n3. The authors need to compare their results with the findings of the Li 2020 paper with synthetic initiation complexes, as\nwell as with the Gao et al paper on initiation complexes of the RNAP-σ54 system (which also has a sigma finger that\nclashes with the 5’end of nascent RNA; https://www.pnas.org/doi/epub/10.1073/pnas.2309670120). Why was no swiveling\nobserved in these structures when the RNA encountered σ3.2?\n4. The authors see a 55-45% split between IC and PIC; what gives rise to this distribution? Further, the ms discusses\ncaptured states; What about dynamic intermediates not captured?\n5. L242: To be able to assess the FRET experiments properly, the FRET assay needs to be properly defined and compared\nwith previous cited work.\n6. L328: There is a suggestion that swiveling causes the DNA scrunching. However, given that scrunching should be\nalready by present in IC6 (caused by the motion of nucleic acids during synthesis of the 6-nt RNA), it appears that swiveling\nis more of a consequence rather than the cause of scrunching.\n7. L399. There are many examples of σ70 retention in E. coli; 2-3 references can be provided. Although the NusG\nhypothesis is a good one, note that in many of these examples, there was no transcription-translation coupling involved (in\nvitro experiments).\nMinor\n1. Annotate the figs better – now hard to link the annotations with the structural elements (e.g., Fig 1b)\n2. L132. Typo (“camp”).\n3. L164. First report of pausing in initiation was in 2016 – so not “for decades”\n4. Ref 51 is incorrect since it refers to a DNA polymerase and a sliding clamp.\n5. L305. Rephrase to clarify that you are refering to swiveling, not only pausing (rather than pausing – since it is long known\nthat σ3.2 is a pausing determinant).\n6. Note that it has been shown that DNA opening in initiation has no obligatory clamp opening/closing\n(https://pubmed.ncbi.nlm.nih.gov/34633286/), so it is best to refrain from talking about RNAP clamp opening/closing (or at\nleast to correct the statement), especially since this is not the conformational change that is the focus on the Zheng ms.\n7. L338: it is unclear why this is considered an allosteric mechanism. In general, it is not clear what the proposed sequence\nof events are as you go from 6 to 7 nt, and then to the paused state.\n8. L379: need to cite Refs 11-13, which established the paused state as a regulatory checkpoint.\n9. L383 and L388: “raised position” and “upward shift” depends on the frame of reference. Provide alternative term, e.g., less-\nengaged state or similar.\n10. Does the 5’ end of RNA has a triphosphate, and if it not, how does it influence the results?", "reviewer_3": "Zheng and Xu report cryo-EM structures of M. tuberculosis (Mtb) RNAP holoenzyme comprising ECF sigE subunit and\ntranscription initiation factor, CarD. Authors solved structures of the paused initiation complex (PIC) with 7-nt RNA and\ninitiation complexes (ICs) with 7- and 6-nt RNAs. Based on the structure analysis, the authors\nsuggest that a clash between sigE region 3.2 and RNA triggers swiveling of the RNAP\nclamp, scrunching of transcription bubble and consequently\ninduces pausing. In addition, they found two conformations of sigE region 4, one of which they attributed to a pre-escape\nstate. Finally they suggest that their results “define a mechanistic checkpoint during transcription initiation” and provide an\nallosteric model for regulation of transcription through RNAP conformational dynamics.\nWhile the cryo-EM data reported here are of good quality and represent a valuable\ncontribution, there is a striking lack of basic biochemical controls to support the\nproposed structures and the conclusions drawn from them. This study has the\npotential to be high-impact, provided the authors supply all necessary data to substantiate their claims. As it stands,\nhowever, the conclusions are not adequately supported by the\navailable data and appear unwarranted.\nThere are two major flaws:\n1. The authors claim to have resolved the structure of a paused complex formed during\ninitial transcription. However, they\nprovide no data demonstrating that an initiation pause actually occurs in their system. In fact, it is unclear how they know\nwhich complex is paused and which is a productive one. While they reference studies\nconducted with E. coli sig70-RNAP,\ntheir work involves an ECF sigma factor and RNAP from Mtb. Mtb RNAP is structurally distinct from E. coli RNAP and\nexhibits different transcriptional behavior. Similarly, ECF sigma factors differ structurally from primary sigma factors, for\nwhich pausing has been previously reported. Thus, E. coli sig70-RNAP pauses at the +6 register, not +7. The authors\nshould perform transcription assays to confirm that initial pausing at the +7 register indeed occurs with the sigE\nRNAP from Mtb.\n2. CarD has been described as a factor regulating the principal sigma subunit,\nparticularly in ribosomal gene transcription.\nWhat is the rationale for including CarD in this study? Is there any published evidence showing that CarD regulates ECF\nsigma-dependent transcription? Is there any data indicating that CarD is involved in initial\npausing? To validate their\nconclusions, the authors should include in vitro transcription assays demonstrating that CarD activates or modulates sigE-\ndependent transcription, influences initial pausing, and stabilizes the sigE-RPo complex.\nSpecific comments:\n1. There is a problem with the experimental setup. First, I found nothing matching the -10 element motif (c/gGTTg/c) for sigE\nrecognition in the sequence provided by authors. To which Mtb promoter corresponds this construct ? That information must\nbe provided. Second, they use template with an artificial bubble of 12 nt. Meantime there are only 10 mismatches alternated\nby two matches. Why ? Is there any reason ?\n2. The authors should cite the original publication that introduces and validates the\nMango transcription assay used.\n3. The FRET experiment (Figure 3d) is neither informative nor conclusive and could\nbe omitted. Control performed with free DNA is required but missing.\n4. page 3 line 47: which “six conserved domains” you mention here ?\n5. page 4 line 50: usually we call sigma 3.2 a region, not a domain.\n6. page 4 line 59: Could the authors clarify what is meant by “simple domains” ?\n7. page 4 line 60: provide a reference to a study demonstrating that the affinity of ECF sigma factors to the RNAP core is\nweaker than that of other sigmas.\n8. page 4 lines 66 – 68: authors wrote that “There is a lack of relevant structural information for Mtb σE factor, it is unclear\nwhether σE factor folds in the same way as other ECF σ factors and adopts a similar strategy to initiate RNAP transcription.”\nThat’s an overstatement. In fact, sigE shares high sequence homology with sigH and recognizes the same promoter motifs.\nThe structure of sigH has been solved, and AlphaFold predicts the structure of sigE with high accuracy.\n9. page 5 line 80: authors wrote “...we investigated the mechanism of pausing during transcription initiation...”\nThere is no data on pausing and therefore on the mechanism of pausing.\n10. page 5 lines 85-86: authors wrote “We revealed that RNA polymerase pauses between transcript lengths of 6 and 7\nnucleotides…”. As I explained above, there is no any data showing that RNAP indeed pauses at +7 register.\n11. page 5 lines 88 – 91: Authors wrote “We also found that CarD binds to non-template ssDNA in the IC complex to\nstabilize the transcription bubble, whereas it interacts with template ssDNA in the PIC complex to stabilize the paused\ntranscriptional conformation.”\nThere is no experimental evidence demonstrating that CarD stabilizes the transcription bubble. If authors wish to prove it,\nthey should perform KMnO4 probing on RPo complexes with and without CarD. Also, there is no data showing that CarD\nstabilizes “paused transcription conformation”. To prove it, authors should perform in vitro transcription kinetics assay with\nand without CarD.\n12. page 6 lines 101-104: authors refer to the studies on E. coli sig70-RNAP reporting pausing at +7 register. This is not\nrelevant because they use a different RNAP and a structurally distinct sigma factor.\n13. page 9 lines 159 – 160: I see no evidence showing that CarD stabilized the non-template ssDNA of the transcription\nbubble.\n14. page 9 lines 163-165: authors wrote “It has been known for decades that RNAP adopts a pausing state when the\nnascent RNA extends to 6 to 7 nucleotides.“\nIf authors refer to the initial transcription pausing by E. coli sig70-RNAP, it has been published in 2016. It’s not “for decades”.\n15. page 11 lines 196-197: authors wrote “… the σ3.2 domain of σE stimulates pausing through its insert into RNA exit\nchannel, forming a roadblock that promotes RNAP swiveling and interrupts nucleotide addition.”\nAs expalined above, there is no biochemical data supporting this conclusion.\n16. page 13 lines 241 – 242: authors wrote “The FRET assay validated this conformational change, consistent with\npreviously reported changes in E. coli. ” The ensemble FRET measurements performed by authors do not include essential\ncontrols which allows to validate it.\n17. page 13 lines 253 – 255: authors wrote “This minimal interface suggests a transient association mechanism at the\ntranscriptional checkpoint, distinct from stable binding.”\nIts not clear what do you mean under “transcriptional checkpoint” here ?\n18. Extended data Figure 1j. The EMSA performed with the bubble template is not informative and adds nothing. It shows\nthat there is binding, but there is nothing to prouve that there is a complex with sigE-CarD-RNAP. RNAP core alone would\nbind and shift this template perfectly well. Also, I do not understand why so huge amounts of RNAP (18 mg) were used ?\nAlso, final concentration of proteins is not mentioned in the Methods section, while it should be.\n19. Extended data Figure 1h . Is it expected that CarD will form a stable complex with sigE (peak2) ? How can it contribute to\nregulation ?\n20. Figure 3. There is a mismatch between Fig.3 panel lettering (panels g, h) and Fig. 3 legend (panels j,k).\n21. EMSA in Figure 3g is not informative. Which DNA was used ? Was there a competitor added ? What is a rational in\ndoing EMSA with sigE alone ?\n22. Figure 4g. A control without CarD is missing. I do not see how this experiment reveals CarD cooperativity .\n23. Overall, manuscript should be rewritten in more concise from. Some sections from Results may go to Introduction (e.g.\np12 lines 222- 227, p14, lines 261-265). Discussion is too excessive. Speculations on NusG and transcription -translation\ncoupling are irrelevant to the current study."}, "author_response": "We are grateful to the Reviewers for their positive comments and critical feedback.\nWe have revised the manuscript in response to their suggestions, addressing all\nquestions and concerns as detailed in the point-by-point responses below.\nIn this revision, we have:\n Clarified the rationale for scaffold design and performed complementary DNA\nscaffold reconstructions, confirming through cryo-EM that the structures\nrepresent physiologically relevant transcription complexes.\n Added new biochemical assays validating the transcriptional pause and testing\nCarD function, which support our structural observations.\n Performed additional controls (DNA-only FRET, EMSAs with defined\ncomponents, RNAP concentrations) and corrected figure legends, labels, and\ntypographical errors.\n Expanded the Methods section to provide full details of cryo-EM data\nprocessing, model building, and validation.\n Revised the Introduction and Discussion for conciseness, integrated selected\nResults into the Introduction, removed speculative text, and clarified\nterminology.\nReferee's remarks are shown in black and our responses are in blue. All modifications\nin the revised manuscript are highlighted in red.\nReviewer #1 (Remarks to the Author):\nThis study reports the three cryo-EM structures of sigma E-RNAP transcription\ninitiation intermediates. There are new insights in the structure PIC7 (paused initiation\ncomplex with 7-nt RNA): (1) a half-translocated state of the DNA-RNA hybrid; (2) a\nswivel conformation of the RNAP resulting in shifted location of upstream dsDNA and\nthe CarD. The author proposed that the steric hindrance between sigma 3.2 and the\nnascent RNA is likely the causing factor inducing pausing at the transcription initiation\nstage. Overall, this study presents an RNAP conformation that has not been observed\nbefore. The flaw in the method compromises the reliability of the conclusion. I have\nfollowing concerns.\nWe thank the reviewer for the positive summary of our work and for highlighting\nthe novelty of the PIC7 structure. We have carefully addressed all the concerns raised,\nas detailed below.\nMajor concerns:\n1. The major concern is the DNA-RNA scaffolds, which have been used to reconstitute\nthe RPitc6 and RPitc7 complexes. The authors did not describe in detail the logic for\ndesigning the DNA-RNA scaffolds. The best way to capture the RPitc6 and RPitc7 is\nby adding subset NTPs to RPo (prepared by incubating RNAP with a fully\ncomplementary promoter DNA) and allowing RNA synthesis to 6 or 7 nt. If the method\nis not feasible, you can reconstitute the complexes with a DNA-RNA scaffold containing\nan RNA of 6 or 7-nucleotide and a double-stranded DNA, but you should use the\ncomplementary sequences for the non-template and template DNA and use an RNA\nwith sequence precisely matching the nascent RNA produced in vivo or in vitro\ntranscription assay. The length of nascent RNA and the size of the transcription bubble\nare closely related. For example, if the RPo has a transcription bubble of 12 bp, then\nan RPitc6 in post-translocation state should have a transcription bubble of 17 bp, and\nan RPitc7 in half-translocated state should also have a transcription bubble of 17\n(RNA has translocated but DNA has not). So if the author would like to use the reconstituted\ncomplexes for structure determination, it is suggested first to identify the transcription\nstart site of a promoter and then to determine an RPo structure with the promoter as\nan unscrunched bubble control. For RPitc6 and RPitc7, the reviewer suggests\ndesigning scaffolds with the correct RNA sequence to define the natural bubble size at\nthe respective states. The current scaffolds have non-complementary sequences in\nthe transcription bubble and incorrect bubble size for RPitc6 and 7, and therefore, they\nare not the physiological-relevant transcription intermediate complexes.\nWe sincerely thank the reviewer for raising this important point regarding the\ndesign of DNA-RNA scaffolds for RPitc6 and RPitc7. We completely agree with the\nReviewer’s opinion and recognize that fully complementary scaffolds are theoretically\nideal for defining natural transcription bubble sizes. We attempted to reconstitute\nRPitc6 and RPitc7 with fully matched template and non-template DNA. At RNA length\n6 nt, RNAP did not pause, and at RNA length 7 nt, RNAP exhibited swivel rotation\nconsistent with pausing. Cryo-EM density for these fully complementary scaffolds was\nweak and partially missing, likely due to reannealing or conformational heterogeneity\n(Extended Data Fig. 4 and Extended Data Fig. 5), indicating technical challenges in\ncapturing these transient intermediates.\nIn previous bacterial RNAP initiation studies (Li et al., PNAS 2020, 117: 5801-\n5809; Li et al., Nat Commun 2019, 10: 1-14; Fang et al., NAR 2019, 47: 7094-7104;\nSupplementary Table S1), pre-melted scaffolds are used to visualize nascent RNA-\nσ3.2 clashes and initiation intermediates. Our design of a ~12 bp transcription bubble\naligns with these precedents, while fully complementary scaffolds are primarily used\nfor RPo. In currently resolved structures of Mycobacterium tuberculosis and\nMycobacterium smegmatis fully complementary scaffolds (Supplementary Table S1)\noften result in loss of transcription bubble density, likely due to the technical challenges\nof this approach. Therefore, we employed non-complementary DNA scaffolds to\ncapture paused conformations during transcription initiation.\nWe also performed in vitro transcription assays with Mtb RNAP, which confirmed\nthat RNAP pauses at RNA lengths of 6-7 nt (Extended Data Fig. 1H, I), validating that\nthe intermediates captured by our pre-melted scaffolds are functional. Together, these\nobservations support that our scaffold design is both experimentally validated and\nconsistent with established structural studies. We agree with the Reviewer that using\nfully complementary scaffolds would in principle be preferable; however, such scaffolds\nfrequently resulted in missing transcription bubble density (Extended Data Fig. 4-5).\nTo overcome this technical limitation and to visualize the intermediates with greater\nstructural detail, we therefore employed non-complementary scaffolds. In the revised\nmanuscript, we have also expanded the sample assembly section to make our scaffold\ndesign logic more explicit (Page 7, Lines 131-135; Page 8, Lines 138-142; Page 9,\nLines 169-181).\n2.The transcription initiation complex with a 7-nt RNA yield two major conformation (IC\nand PIC). The authors should give some explanation for why these two distinct\nconformations arise. Do these conformations represent sequential intermediates, or\nare they alternative pathways during transcription?\nWe thank the reviewer for these insightful questions. The synthesis of 6-7 nt RNAs\nrepresents a transcriptional checkpoint where RNAP frequently pauses and may\nsample multiple outcomes, including promoter escape or abortive release (e.g.,\nKapanidis et al., Science 2006, 314: 1144-1147; Duchi et al., Mol Cell 2016, 63: 939-\n950). Consistent with this, our cryo-EM analysis yielded two major conformations (IC\nand PIC) that likely reflect different states sampled at this checkpoint.\nWe apologize that our study cannot definitively resolve whether IC and PIC\nrepresent sequential intermediates along a single pathway or alternative\nconformational outcomes. Current knowledge in the field also does not provide a\ndefinitive answer. Our data demonstrate coexistence of both states but cannot by\nthemselves resolve their temporal relationship. As there is no consensus in the field,\nwe have revised the text to describe them as distinct conformations observed at the 7-\nnt checkpoint, without assuming a defined order or pathway. In the revised manuscript,\nwe have emphasized this “checkpoint” view in the Results (Page 8, Lines 152-158;\nPage 16, Lines 309-312) and Discussion (Page 22, Lines 448-450), highlighting that\npausing at 6-7 nt represents a branching point that influences the outcome of\ntranscription initiation.\n3.Figure 3a,b-The authors should include zoomed-in views to highlight the contrasting\nregions of ‘unwinding’ and ‘non-unwound’ template DNA. The current version does not\nshow this with enough clarity. The DNA should be labeled relative to the TSS.\nWe thank the reviewer for this helpful suggestion to improve the clarity of Figure\n3A and B. In the revised version, we have added zoomed-in panels to highlight the\ncontrasting regions of unwound versus non-unwound template DNA in both complexes,\nproviding clearer visual contrast. The template DNA positions are now labeled relative\nto the transcription start site (TSS), allowing the register of unwinding to be directly\nappreciated. We have also updated the figure legend to explicitly describe the\nhighlighted regions and the TSS reference. These revisions, now shown in Figure 3A\nand B, provide a clearer comparison of the unwound and non-unwound DNA segments.\n4. The manuscript lacks a clear explanation of how the authors selected the particle\nclasses of interest during cryo-EM data processing. It is recommended to clarify\nwhether selection was based on resolution, particle percentages, or other criteria.\nPlease provide this information either in the Extended Data Figures or in the Methods\nsection.\nWe thank the Reviewer for critical questions and insightful suggestions. During\ncryo-EM data processing, particle classes were selected based on the presence of\ncomplete complexes, rather than on resolution or particle percentages alone.\nSpecifically, we retained only those classes that clearly contained all of the expected\ncomponents of the transcription initiation assembly, including the RNAP core, σE, CarD,\nand the nucleic acid scaffold.\nThis criterion ensured that the analyzed particles represented intact initiation\ncomplexes suitable for structural interpretation. To clarify this point, we have revised\nthe Methods section (Page 31, Lines 632-634), and added explanatory notes in the\nExtended Data Figures 2-5 showing the classification workflow.\nMinor issues:\n1. Extended Data Fig. 2: The label “Mtb IC6” appears to be incorrect and should be\nrevised to “Mtb IC7”.\nWe thank the reviewer for catching this labeling error and apologize for the\noversight. We agree the label in Extended Data Fig. 2 should read “Mtb IC7” (not\n“Mtb IC6”). We have corrected the panel label and updated the figure legend (Page\n45, Lines 785) and all in-text references to ensure consistency across the manuscript.\nWe also re-checked all figure and Extended Data labels for IC/PIC and 6-7 nt\ndesignations to avoid similar inconsistencies elsewhere.\n2. More detail is needed regarding the model building procedure, particularly for how\nthe DNA and protein components were modeled and validated.\nWe thank the Reviewer for these critical questions and insightful suggestions. In\nthe revised manuscript, we have expanded the Methods section to provide more detail\non model building. The models for RNAP, σE, CarD, and the nucleic acid scaffold were\ninitially based on previously reported structures (PDB 5ZX2, 6EDT, 6JBQ). Each\ncomponent was rigid-body fitted into the cryo-EM maps using Chimera, followed by\nlocal adjustments through flexible fitting in COOT. The DNA/RNA scaffold was\nmanually built into the density guided by the cryo-EM map and prior scaffold designs\n(Fang et al., NAR 2019, 47: 7094-7104), with RNA positioning adjusted to match\nobserved density, particularly in the σ3.2 region. The complete models were refined in\nPhenix using real-space refinement, with secondary structure, geometry, and non-\ncrystallographic symmetry restraints applied. Model geometry was assessed using\nMolProbity, and map-to-model fit was evaluated by FSC between the model and the\nmap. The DNA and RNA densities are well-resolved in the analyzed regions,\nconfirming the accuracy of nucleic acid positioning. These details have been\nincorporated into the Methods section (Page 32, Lines 649-658) in the revised\nmanuscript.\n3. Line 474: “a 6 polyacrylamide native gel” should be “a 6% polyacrylamide native\ngel”.\nWe thank the reviewer for pointing out this typographical error and apologize for\nthe oversight. The manuscript has been updated so that “a 6 polyacrylamide native gel”\nnow correctly reads “a 6% polyacrylamide native gel” in the Methods section (Page 27,\nLines 544-545).\n4. Figure 2d: The DNA template nucleotides in the active site appear to be missing.\nWe thank the reviewer for pointing this out and apologize for the oversight in the\nfigure preparation. The apparent absence of DNA template nucleotides in the active\nsite was due to the transparency settings used during figure preparation. We have re-\nrendered Figure 2d with adjusted visualization settings to clearly show the DNA\ntemplate nucleotides in the RNAP active site.\nReviewer #2 (Remarks to the Author):\nThe work of Zheng et al provides structural snapshots of important intermediates of\nthe transcription path from initiation to elongation, centred around the pause site that\nhas been observed to occur after the synthesis of around 6-7 nt of RNA. Despite the\navailable information biochemical and biophysical information, there are no high-\nresolution structures of the paused state. Zheng et al used cryoEM to provide such\ninformation for the Mtb system and reveal several interesting conformational changes,\nthe most exciting being the presence of a half-translocated state with RNAP swiveling\nin the paused complex (which to my knowledge has not been observed before in\ninitiation), a motion that has been associated with different types of pausing in\ntranscription elongation. This observation draw nice parallels between initiation and\nelongation and offers an additional element for the regulation of transcription initiation\nin bacteria, which may have generality beyond the Mtb system.\nThe paper is overall well written, although there are points that needs attention, and\naspects that needs addressing before publication.\nWe thank the reviewer for the thoughtful summary and for highlighting the novelty\nof the half-translocated and swiveling states in the paused initiation complex. We have\naddressed all specific points raised in detail below.\nMajor points\n1. The authors need to be much more explicit about the fact that the complexes\nprepared are synthetic, i.e., not prepared during actual synthesis, but assembled using\nsynthetic nucleic acid scaffolds (Ext Fig 4) – as a result, it appears that the complexes\nhave much less scrunching that actual complexes with the same RNA length would\nhave had, and do not reflect exactly the final structures of complexes formed via actual\nRNA synthesis. For example, for the template provided, an IC7 would expect to melt\nthe region from +1 to +7; however, currently the RNA is hybridized using the existing\nbubble. The authors need to discuss how their main and secondary findings are\naffected by the assembly method, and why this assembly route was pursued.\nWe thank the Reviewer for raising this important point and acknowledge that our\ninitiation complexes were assembled using synthetic nucleic acid scaffolds rather than\nbeing formed during active RNA synthesis. Consequently, the extent of scrunching may\nbe less than in fully transcribing complexes of the same RNA length, and the RNA-\nDNA hybrid is constrained by the pre-formed bubble.\nWe did attempt to assemble complexes with fully complementary template and\nnon-template DNA, but the cryo-EM density was weaker and partially missing, likely\ndue to reannealing or increased conformational heterogeneity (Extended Data Fig. 4\nand Extended Data Fig. 5). Although pre-melted scaffolds indeed reduce the degree\nof scrunching compared with fully transcribing complexes, this approach enables\nstable capture of paused intermediates suitable for structural analysis. Based on\nprevious bacterial RNAP initiation studies (Li et al., PNAS 2020, 117: 5801-5809; Li et\nal., Nat Commun 2019, 10: 1-14; Fang et al., NAR 2019, 47: 7094-7104 ；\nSupplementary Table S1), using pre-melted transcription bubbles is a well-\nestablished strategy to visualize nascent RNA-σ3.2 clashes, and our ~12-bp bubble\ndesign is consistent with these precedents. In the revised manuscript, we have also\nexpanded the sample assembly section to make our scaffold design logic more explicit\n(Page 7, Lines 131-135; Page 8, Lines 138-142; Page 9, Lines 169-181). We also\nperformed in vitro transcription assays, which confirmed that Mtb RNAP pauses at RNA\nlengths of 6-7 nt (Extended Data Fig. 1H, I), validating that the intermediates captured\nby our scaffolds are functional.\nWe note that while pre-melted scaffolds may exhibit less scrunching than fully\ntranscribing complexes, the captured intermediates still represent functional relevant\nstates. In particular, we observed RNAP swivel conformations associated with\ntranscriptional pausing, which directly link RNAP structural rearrangements to pause\nformation. These findings are consistent with the results obtained using non-\ncomplementary scaffolds and do not affect our analysis of RNAP rotation and pausing.\nAlthough pre-melted DNA may slightly alter the degree of transcription bubble\nscrunching, this does not impact the core conclusions of our study. We have now\nexplicitly discussed how the main and secondary findings are affected by the assembly\nmethod in the Discussion (Page 18, Lines 364-370).\n2. L62. Do we know that the picture for the ECF σE is similar as for σA? Why was σΑ\nnot selected to proceed with structural analysis?\nWe thank the reviewer for this thoughtful question. We agree that the overall\npicture for σE is similar to that for σA. we note that some structural studies of σA-\ndependent initiation complexes have not captured the swiveling conformations\nassociated with pausing, likely because σA contains both the σ3 domain and a flexible\nσ3.2 linker, which together may restrict the overall conformational flexibility of the\ncomplex. By contrast, σE possesses only a flexible σ3.2 linker without the additional\nσ3 domain, resulting in a simpler domain organization and conformational plasticity. (Li\net al., PNAS 2020, 117: 5801-5809; Pukhrambam et al. ， PNAS 2022, 119:\ne2201301119). For this reason, we chose σE, which has a simpler domain organization\nand greater conformational plasticity, making it more suitable for resolving dynamic\npaused states and RNA-σ3.2 interactions. The reduced modules complexity of σE (σ2,\nσ3.2, σ4) compared to σA (σ1.1, σ1.2, σ2, σ3, σ3.2, σ4) also minimizes conformational\nheterogeneity, facilitating high-resolution structural analysis. We have clarified this\nrationale in the Introduction of the revised manuscript (Page 4, Lines 65-73).\n3. The authors need to compare their results with the findings of the Li 2020 paper with\nsynthetic initiation complexes, as well as with the Gao et al paper on initiation\ncomplexes of the RNAP-σ54 system (which also has a sigma finger that clashes with\nthe 5’end of nascent RNA; https://www.pnas.org/doi/epub/10.1073/pnas.2309670120). Why was no swiveling\nobserved in these structures when the RNA encountered σ3.2?\nWe thank the Reviewer for these critical and insightful suggestions. In the Li et al.\n(Li et al., PNAS 2020, 117: 5801-5809) study, the DNA scaffold lacked the upstream\n−35 region and did not contain a fully formed transcription bubble, likely prevented\ndetection of large-scale RNAP conformational changes. In our cryo-EM analysis with\nσE, the inclusion of upstream promoter elements and a complete bubble scaffold\nallowed us to visualize the swiveling motion in coordination with RNA-σ3.2 interactions.\nIn the Gao et al. (Gao et al., PNAS 2024, 121: e2309670120) study, the E. coli σ54\nsystem also features a σ finger that clashes with the 5′ end of RNA, but σ54 has distinct\nstructural features and regulatory mechanisms compared to the mycobacterial σE-\nRNAP system, which likely explains why swiveling was not observed. We have added\nthese comparisons in the Discussion section (Page 20, Lines 395-402).\n4. The authors see a 55-45% split between IC and PIC; what gives rise to this\ndistribution? Further, the ms discusses captured states; What about dynamic\nintermediates not captured?\nWe thank the reviewer for this insightful question. In the revised manuscript, we\nhave added a sentence in the Results section (Page 8, Lines 152-158) to clarify the\nobserved ~55:45% distribution between IC and PIC. This distribution likely reflects the\nrelative stability and population of these two conformations under the experimental\nconditions, representing alternative paused states within the initiation checkpoint.\nWe also acknowledge that additional dynamic intermediates likely exist, including\npartially scrunched states, transiently backtracked complexes, or short-lived RNA-σ3.2\ninteractions, which were not captured due to their low abundance and rapid transitions.\nThese limitations are inherent to cryo-EM, which favors the visualization of more\npopulated and stable states. We have explicitly discussed the possibility of such\ndynamic intermediates in the Results section (Page 8, Lines 152-158).\n5. L242: To be able to assess the FRET experiments properly, the FRET assay needs\nto be properly defined and compared with previous cited work.\nWe thank the Reviewer for critical questions and insightful suggestions. We have\nnow clarified the description of our FRET assay in the Results section (Page 15, Lines\n293-298), emphasizing how our bulk FRET design (with Cy3/Cy5 labeling) captures\nRNA length-dependent conformational changes. We also compared our findings with\npreviously reported sm-FRET studies, highlighting that both approaches detect a clear\nsignal shift when RNA extends from 6 to 7 nt (Page 15, Lines 293-298).\n6. L328: There is a suggestion that swiveling causes the DNA scrunching. However,\ngiven that scrunching should be already by present in IC6 (caused by the motion of\nnucleic acids during synthesis of the 6-nt RNA), it appears that swiveling is more of a\nconsequence rather than the cause of scrunching.\nWe sincerely thank the reviewer for this insightful and important observation. We\nfully agree with the reviewer’s point that scrunching is already present in IC6 due to\nRNA synthesis, and that swiveling should not be regarded as its primary cause. This\nis a critical mechanistic insight, and we greatly appreciate the reviewer highlighting it.\nOur structural data are consistent with this interpretation: swiveling becomes more\npronounced in PIC7, where it appears to further stabilize the scrunched bubble and\nreinforce the paused state rather than to initiate scrunching. We have updated the\nDiscussion (Page 20, Lines 389-392) to make this distinction explicit and to note that\nresolving the temporal relationship between scrunching and swiveling is an important\nkinetic question for future investigations.\n7. L399. There are many examples of σ70 retention in E. coli; 2-3 references can be\nprovided. Although the NusG hypothesis is a good one, note that in many of these\nexamples, there was no transcription-translation coupling involved (in vitro\nexperiments).\nWe thank the reviewer for the comment on σ factor retention. We have added\nthree references on σ70 retention in E. coli (Reference 68-70) to provide context and\nhighlight that σ factor retention during elongation is well-documented in E. coli. We\nthank the reviewer for the comment regarding NusG. We agree that, as noted, there is\nno direct evidence linking NusG to σE retention in our system; our previous discussion\nwas speculative. Therefore, following the reviewer’s suggestion that this discussion is\nnot directly relevant, we have removed this portion from the manuscript.\nMinor\n1. Annotate the figs better – now hard to link the annotations with the structural\nelements (e.g., Fig 1b)\nWe thank the reviewer for this suggestion. We have improved the figure\nannotations in all relevant panels (e.g., Fig. 1B, E) to clearly link labels with the\ncorresponding structural elements. The updated figures now include more precise\npointers and consistent color coding to enhance clarity.\n2. L132. Typo (“camp”).\nWe thank the reviewer for pointing out this typographical error and apologize for\nthe oversight. The typo “camp” on line 132 has been corrected in the revised\nmanuscript (Page 10, Line 187).\n3. L164. First report of pausing in initiation was in 2016 – so not “for decades”\nWe thank the reviewer for the correction and apologize for the imprecise statement\nin the original manuscript. We have revised the text to accurately reflect that the first\nreport of transcriptional pausing during initiation was in 2016, and removed the phrase\n“for decades” (Page 11, Line 218-220).\n4. Ref 51 is incorrect since it refers to a DNA polymerase and a sliding clamp.\nWe thank the reviewer for pointing this out and apologize for the incorrect\nreference in the original manuscript. Reference 51 has been corrected and is now\nlisted as Reference 55. We have updated the citation to (Robb et al., 2013 JMB, 425:\n875-885; Bae et al., 2015 eLife, 4: e08504), which provide structural and sm-FRET\nevidence for open promoter complexes (RPo) with unwound transcription bubbles of\n~12-14 bp.\n5. L305. Rephrase to clarify that you are refering to swiveling, not only pausing (rather\nthan pausing – since it is long known that σ3.2 is a pausing determinant).\nWe thank the Reviewer for critical questions and insightful suggestions. The\nsentence previously on line 305 has been rephrased in the revised manuscript (Page\n18, Lines 360-362) to clarify that we are referring to swiveling, not only pausing, and\nnow explicitly distinguish the role of σ3.2 in swiveling from the pausing observed in our\nstructures.\n6. Note that it has been shown that DNA opening in initiation has no obligatory clamp\nopening/closing (https://pubmed.ncbi.nlm.nih.gov/34633286/), so it is best to refrain\nfrom talking about RNAP clamp opening/closing (or at least to correct the statement),\nespecially since this is not the conformational change that is the focus on the Zheng\nms.\nWe thank the Reviewer for this important clarification, and we apologize that the\nwording in our previous manuscript was not rigorous. The manuscript has been revised\n(Page 19, Lines 377–379) to avoid implying that RNAP clamp opening/closing is\nrequired for DNA opening during initiation, and instead focus on the conformational\nchanges highlighted in our study. The statement has been corrected accordingly.\n7. L338: it is unclear why this is considered an allosteric mechanism. In general, it is\nnot clear what the proposed sequence of events are as you go from 6 to 7 nt, and then\nto the paused state.\nWe thank the Reviewer for this insightful comment. In our structural analysis, we\nobserved that when the RNA reaches 6 nt, RNAP does not enter the paused state.\nHowever, upon extension to 7 nt, RNAP undergoes a conformational change—\nspecifically, swiveling of certain modules—leading to transcriptional pausing. This\nchange is triggered by a steric clash between the 5′ end of the nascent RNA and the\nσ3.2 region. Based on these observations, we propose that the pausing mechanism of\nRNAP represents an RNA-induced allosteric process occurring during the transition\nfrom 6 to 7 nt. We performed in vitro transcription assays, which confirm that RNAP\npauses as RNA extends from 6 to 7 nt. In the revised manuscript, we have clarified\nthat the term “allosteric” refers to RNA-induced conformational changes in RNAP,\noutlined the proposed sequence of events leading to the paused state, and included\nthe experimental evidence validating the pausing at the 6 nt to 7 nt. We also note that\nwhile this model provides a plausible mechanistic explanation, the precise sequence\nand causal relationships remain to be fully established, and these points have been\nincorporated into the Discussion (Page 20, Lines 403–409).\n8. L379: need to cite Refs 11-13, which established the paused state as a regulatory\ncheckpoint.\nWe thank the reviewer for pointing this out, and we apologize that the wording in\nour previous manuscript was not rigorous. We have now cited References 11-13 to\nhighlight that the paused state functions as a regulatory checkpoint during transcription\ninitiation (Page 22, Lines 446).\n9. L383 and L388: “raised position” and “upward shift” depends on the frame of\nreference. Provide alternative term, e.g., less-engaged state or similar.\nWe thank the reviewer for this helpful suggestion. To avoid frame-dependent\nterminology, we have revised (Page 23, Lines 451; 461–465) the text to replace\n“raised position” and “upward shift” with more neutral descriptions such as “less-\nengaged state” and “reduced engagement”.\n10. Does the 5’ end of RNA has a triphosphate, and if it not, how does it influence the\nresults?\nWe thank the reviewer for this perceptive and important point — the chemical state\nof the RNA 5′ end is indeed relevant to interpreting pausing. If the nascent RNA lacked\na 5′-triphosphate, the steric interaction between the 5′ end and the σ3.2 tip would likely\nbe weakened; this could reduce the dwell time of the +7 pause (and in extreme cases\nalter the apparent register of the pause), so the absence of a 5′-triphosphate could\nmeaningfully affect the biochemical readout and its structural interpretation. To avoid\nthis potential confound, all RNAs used in our experiments carried a natural 5′-\ntriphosphate, which we have explicitly noted in the Methods (Page 26, Lines 518; 529).\nIn the cryo-EM maps, the triphosphate group is clearly resolved in IC6 but not in IC7\nand PIC7, most likely due to increased flexibility of the 5′ end as the RNA extends.\nImportantly, the use of 5′-triphosphate RNA ensures that our biochemical and structural\nanalyses reflect the physiological state and do not affect our conclusions on the paused\nstates and the RNA-σ3.2 interactions.\nReviewer #3 (Remarks to the Author):\nZheng and Xu report cryo-EM structures of M. tuberculosis (Mtb) RNAP holoenzyme\ncomprising ECF sigE subunit and transcription initiation factor, CarD. Authors solved\nstructures of the paused initiation complex (PIC) with 7-nt RNA and transcription\ninitiation factor, CarD. Authors solved structures of the paused initiation complex (PIC) with 7-nt RNA and\ninitiation complexes (ICs) with 7- and 6-nt RNAs. Based on the structure analysis, the authors\nsuggest that a clash between sigE region 3.2 and RNA triggers swiveling of the RNAP\nclamp, scrunching of transcription bubble and consequently\ninduces pausing. In addition, they found two conformations of sigE region 4, one of which they attributed to a pre-escape\nstate. Finally they suggest that their results “define a mechanistic checkpoint during transcription initiation” and provide an\nallosteric model for regulation of transcription through RNAP conformational dynamics.\nWhile the cryo-EM data reported here are of good quality and represent a valuable\ncontribution, there is a striking lack of basic biochemical controls to support the\nproposed structures and the conclusions drawn from them. This study has the\npotential to be high-impact, provided the authors supply all necessary data to substantiate their claims. As it stands,\nhowever, the conclusions are not adequately supported by the\navailable data and appear unwarranted.\nWe thank the reviewer for the constructive summary of our work and for\nhighlighting the potential impact of our study. We have carefully addressed all the\nspecific concerns raised, as detailed below.\nThere are two major flaws:\n1. The authors claim to have resolved the structure of a paused complex formed during\ninitial transcription. However, they\nprovide no data demonstrating that an initiation pause actually occurs in their system. In fact, it is unclear how they know\nwhich complex is paused and which is a productive one. While they reference studies\nconducted with E. coli sig70-RNAP,\ntheir work involves an ECF sigma factor and RNAP from Mtb. Mtb RNAP is structurally distinct from E. coli RNAP and\nexhibits different transcriptional behavior. Similarly, ECF sigma factors differ structurally from primary sigma factors, for\nwhich pausing has been previously reported. Thus, E. coli sig70-RNAP pauses at the +6 register, not +7. The authors\nshould perform transcription assays to confirm that initial pausing at the +7 register indeed occurs with the sigE\nRNAP from Mtb.\nWe thank the reviewer for this important comment and apologize that the original\nmanuscript did not include in vitro transcription data to directly demonstrate initial\ntranscription pausing in the Mtb RNAP-σE system. Our structural analysis shows that\nwhen the nascent RNA extends from 6 to 7 nt, RNAP undergoes a swivel rotation,\nwhich distorts the bridge helix (BH) and stabilizes a paused transcription complex. This\nstructural feature allows us to assign PIC7 as a paused state. We also recognize that\nMtb RNAP and ECF σ factors differ structurally and functionally from E. coli RNAP and\nprimary σ factors, as the reviewer noted.\nTo directly address this concern, we performed in vitro transcription assays with\nMtb RNAP-σE using a fluorescence-based labeling approach (Wang et al., 2025\nNature, 643: 1127-1134). These assays show two distinct RNA species corresponding\nto 6-nt and 7-nt transcripts (Extended Data Fig. 1H, I). The 6-nt band is more\nprominent, while the 7-nt band is weaker, consistent with previous observations in E.\ncoli σ70-RNAP by Duchi et al., Mol Cell 2016, 63: 939–950. These data indicate that\npausing occurs during the transition from 6-nt to 7-nt RNA, with the 6-nt species\nrepresenting a post-translocation, pre-paused intermediate, and the 7-nt species\nrepresenting the paused state (Page 7, Lines 131-135).\nTogether, the cryo-EM structures and the biochemical data validate the\nassignment of PIC7 as a functional relevant paused state in Mtb RNAP-σE transcription\ninitiation.\n2. CarD has been described as a factor regulating the principal sigma subunit,\nparticularly in ribosomal gene transcription. What is the rationale for including CarD in\nthis study? Is there any published evidence showing that CarD regulates ECF sigma-\ndependent transcription? Is there any data indicating that CarD is involved in initial\npausing? To validate their\nconclusions, the authors should include in vitro transcription assays demonstrating that CarD activates or modulates sigE-\ndependent transcription, influences initial pausing, and stabilizes the sigE-RPo complex.\nWe thank the reviewer for raising this important question. We agree with that CarD\nhas been well described as a factor regulating the principal sigma subunit, particularly\nin ribosomal gene transcription. We noted that in Myxococcus xanthus, CarD can\nmodulate ECF sigma factor activity (Abellón-Ruiz et al., Environ Microbiol. 2014,\n16:2475-2490). Since M. xanthus and M. tuberculosis are both actinobacteria and\nCarD shows high sequence homology between them, this finding suggests that its\nregulatory role may be conserved. Based on this evidence, we included CarD in our\nstudy of Mtb σE-dependent transcription. In the revised manuscript, we have added\nthis reference to highlight the rationale for including CarD (References 46) and also\nexpanded the Introduction to highlight the rationale for investigating CarD in the σE\nsystem (Page 5, Lines 89–93).\nTo directly address the reviewer’s question regarding whether CarD is involved in\ninitial pausing, we performed in vitro transcription experiments (Extended Data Fig.\n1H, I). These assays show that CarD promotes escape from initial transcriptional\npausing: the intensity of the paused 7-nt RNA band is reduced upon addition of CarD.\nThis suggests that CarD assists in pausing escape, likely by interacting with the non-\ntemplate strand to stabilize the transcription complex and facilitate transcription bubble\nopening.\nFurthermore, FRET-based experiments were conducted to test the stabilization of\nthe RPo complex. As shown in Fig. 4G, CarD increases the stability of the σE-RNAP-\nDNA complex, consistent with its proposed role in facilitating productive transcription\ninitiation. Together, these results provide experimental evidence supporting the\nregulatory role of CarD in ECF σE-dependent transcription in Mtb.\nSpecific comments:\n1. There is a problem with the experimental setup. First, I found nothing matching the\n-10 element motif (c/gGTTg/c) for sigE recognition in the sequence provided by authors.\nTo which Mtb promoter corresponds this construct ? That information must be provided.\nSecond, they use template with an artificial bubble of 12 nt. Meantime there are only\n10 mismatches alternated by two matches. Why ? Is there any reason ?\nWe thank the reviewer for raising these important points and sincerely apologize\nthat in our initial scaffold design we did not fully recognize these issues. First, regarding\nthe promoter sequence, we acknowledge that the scaffold we used did not contain the\ncanonical -10 element motif (c/gGTTg/c) for σE recognition. Our original design\nfollowed the cryo-EM study (Fang et al., NAR 2019, 47: 7094-7104), which employed\na pre-melted scaffold to capture a chimeric Mtb σH/σE complex. For the artificial\ntranscription bubble, we similarly followed the design (Fang et al., NAR 2019, 47: 7094-\n7104), which contained 12 nt with 10 mismatches interspersed by two matched bases.\nIn our structures, these two bases are nevertheless fully unwound, suggesting that\ntheir presence does not influence the bubble architecture or the interactions relevant\nto pausing (Page 8, Lines 138-142; Page 9, Lines 169-181).\nWe very much agree with the reviewer that addressing these issues is essential\nfor strengthening our conclusions. Therefore, we redesigned scaffolds to incorporate\na canonical -10 motif and performed in vitro transcription assays, which confirmed\npausing at the +7 position (Extended Data Fig. 1H, I). We also tested fully\ncomplementary DNA scaffolds to capture the PIC7 state (Page 9, Lines 169-181).\nAlthough the density was weaker, likely due to scaffold instability or the transient nature\nof this intermediate, the overall conformation and swiveling observed were consistent\nwith the scaffold-assembled complexes. Together, these additional experiments\nsupport that the structures we report represent the paused state (Extended Data Fig.\n4 and Extended Data Fig. 5).\n2. The authors should cite the original publication that introduces and validates the\nMango transcription assay used.\nWe thank the reviewer for the suggestion, and we apologize that the wording in\nour previous manuscript was not rigorous. The Mango transcription assay used in our\nstudy was first introduced and validated by Dolgosheina et al., ACS Chem Biol 2014,\n9: 2412-2420. We have now added this citation in the Methods section to properly\nacknowledge the original publication (Page 30, Lines 597).\n3. The FRET experiment (Figure 3d) is neither informative nor conclusive and could\nbe omitted. Control performed with free DNA is required but missing.\nWe thank the Reviewer for critical questions and insightful suggestions. We agree\nthat the FRET experiment provides limited mechanistic insight. Therefore, we have\nmoved this experiment to Extended Data Fig. 7F rather than the main figure. However,\nit serves to confirm that our scaffold design is consistent with previous studies. As a\ncontrol, we have now included measurements with free DNA, showing negligible FRET\nsignal, which supports that the observed signal in Extended Data Fig. 7F arises from\nthe assembled transcription complex. We have clarified this in the revised figure legend\n(Page 54, Lines 863–864).\n4. page 3 line 47: which “six conserved domains” you mention here ?\nWe thank the reviewer for pointing this out, and we apologize that the wording in\nour previous manuscript was not rigorous. The phrase “six conserved domains” was\ninaccurate. We have corrected the text to refer to “modules” instead, specifying that σE\ncontains only σ2, σ4, and σ3.2 modules, whereas σA contains σ1.1, σ1.2, σ2, σ3, σ3.2,\nand σA contains σ1.1, σ1.2, σ2, σ3, σ3.2,\nand σ4 modules. This more accurately reflects the structural organization of these σ\nfactors. We have corrected the manuscript accordingly (Page 3, Lines 48–52).\n5. page 4 line 50: usually we call sigma 3.2 a region, not a domain.\nWe thank the reviewer for the comment, and we apologize that the wording in our\nprevious manuscript was not rigorous. We have revised the manuscript to refer to σ3.2\nas a “region” rather than a “domain,” in line with the standard terminology (Page 3,\nLines 48–52). We have corrected the manuscript accordingly.\n6. page 4 line 59: Could the authors clarify what is meant by “simple domains” ?\nWe thank the reviewer for pointing this out, and we apologize that the wording in\nour previous manuscript was not rigorous. We have replaced “simple domains” with\n“structurally simpler regions” to avoid confusion and better reflect the architecture of\nσE compared to σA. We have corrected the manuscript accordingly (Page 4, Lines 66).\n7. page 4 line 60: provide a reference to a study demonstrating that the affinity of ECF\nsigma factors to the RNAP core is weaker than that of other sigmas.\nWe thank the reviewer for this valuable comment and apologize that we could not\nidentify a study directly demonstrating that ECF σ factors have the weakest affinity\namong all σ families. We have now cited the study by Maeda et al. (Nucleic Acids Res.\n2000), which systematically compared the binding affinities of seven E. coli σ factors\nto the RNAP core. This study showed that σE along with other alternative σ factors,\nbinds core RNAP weaker than the primary σ70/σA. In the revised manuscript, we have\ntherefore phrased this point more precisely as “reduced RNAP-binding stability are the\nreasons we selected σE rather than σA for our study.” (Page 4, Lines 66–68)\n8. page 4 lines 66 – 68: authors wrote that “There is a lack of relevant structural information for Mtb σE factor, it is unclear\nwhether σE factor folds in the same way as other ECF σ factors and adopts a similar strategy to initiate RNAP transcription.”\nThat’s an overstatement. In fact, sigE shares high sequence homology with sigH and recognizes the same promoter motifs.\nThe structure of sigH has been solved, and AlphaFold predicts the structure of sigE with high accuracy.\nWe thank the reviewer for pointing this out, and we apologize that the wording in\nour previous manuscript was not rigorous. We agree that our original statement was\nan overstatement. σE indeed shares high sequence homology with σH and recognizes\nthe same promoter motifs. The structure of σH has been experimentally resolved, and\nAlphaFold predictions indicate that σE adopts a highly similar fold. We have revised the\nmanuscript accordingly to reflect that the structural features of σE are expected to\nclosely resemble those of σH, while still noting that direct experimental validation\nremains valuable (Page 5, Lines 75–77).\n9. page 5 line 80: authors wrote “...we investigated the mechanism of pausing during transcription initiation...”\nThere is no data on pausing and therefore on the mechanism of pausing.\nWe thank the reviewer for this important comment. We acknowledge that in the\noriginal submission we did not provide direct biochemical evidence of pausing.\nStructurally, however, our cryo-EM analysis revealed hallmark features of a paused\nstate: at 7-nt RNA, RNAP undergoes a pronounced swivel movement that distorts the\nbridge helix (BH), a conformational change known to be tightly linked to transcriptional\npausing. These features indicate that the PIC7 complexes represent bona fide paused\nintermediates.\nTo substantiate this interpretation, we have now performed in vitro transcription\nassays using Mtb σE-RNAP on templates containing the canonical -10 element motif.\nThese assays revealed distinct RNA products of 6 and 7 nucleotides (Extended Data\nFig. 1H, I). The 6-nt transcript is more prominent, consistent with a post-translocation\nintermediate preceding pausing, while the weaker 7-nt transcript reflects accumulation\nat the paused state. The weaker intensity of the 7-nt band is expected, since the pause\nis transient and the majority of complexes either remain at 6 nt or proceed into\nelongation.\nTaken together, the combination of structural signatures (swiveling and BH\ndistortion) and functional data (discrete 6-nt and 7-nt RNAs) confirm that our structures\ncapture paused initiation complexes. We have revised the Results section (Page 7,\nLines 131-135) to include these new experimental findings.\n10. page 5 lines 85-86: authors wrote “We revealed that RNA polymerase pauses between transcript lengths of 6 and 7\nnucleotides…”. As I explained above, there is no any data showing that RNAP indeed pauses at +7 register.\nWe appreciate the reviewer’s concern, and we apologize that our previous\ndescription was not sufficiently supported by biochemical evidence. In the revised\nmanuscript, we have now included in vitro transcription assays with Mtb σE-RNAP on\na template containing the -10 element motif (Extended Data Fig. 1H, I). These assays\nshow RNA products corresponding to 6- and 7-nucleotide lengths, with the 6-nt band\nmore prominent and the 7-nt band weaker, consistent with previous observations in E.\ncoli σ70-RNAP by Duchi et al., Mol Cell 2016, 63: 939–950. These data indicate that\npausing occurs during the transition from 6-nt to 7-nt RNA, with the 6-nt species\nrepresenting a post-translocation, pre-paused intermediate, and the 7-nt species\nrepresenting the paused state. Our structural data also reveal that RNAP undergoes a\nconformational change associated with pausing when the RNA extends from 6 to 7\nnucleotides. Together, the structural and biochemical data provide support for our\nconclusion that RNAP pauses at the 6 to 7 nt register in this system.\n11. page 5 lines 88 – 91: Authors wrote “We also found that CarD binds to non-template ssDNA in the IC complex to\nstabilize the transcription bubble, whereas it interacts with template ssDNA in the PIC complex to stabilize the paused\ntranscriptional conformation.”\nThere is no experimental evidence demonstrating that CarD stabilizes the transcription bubble. If authors wish to prove it,\nthey should perform KMnO4 probing on RPo complexes with and without CarD. Also, there is no data showing that CarD\nstabilizes “paused transcription conformation”. To prove it, authors should perform in vitro transcription kinetics assay with\nand without CarD.\nWe thank the reviewer for pointing out that there is no direct experimental\nevidence demonstrating that CarD stabilizes the transcription bubble. We apologize\nthat our previous description was not sufficiently rigorous. To address this, we\nperformed FRET experiments using fluorescently labeled transcription bubble\nscaffolds (following Cordes et al., Biochemistry 2010, 49:9171-9180), which show that\nCarD promotes bubble stability in the σE-RNAP system (Fig. 4G). Although KMnO₄\nprobing was suggested, we were unable to perform this assay due to restrictions on\npiperidine in China. The FRET approach serves as a widely used alternative to monitor\ntranscription bubble dynamics.\nWe also thank the reviewer for raising the concern regarding the statement that\nCarD stabilizes the “paused transcription conformation.” We apologize that this\ndescription was imprecise. Based on our structural and biochemical data, a more\naccurate interpretation is that in the initiation complex (IC), CarD interacts with the\ntemplate strand to maintain the open complex. In the paused initiation complex (PIC),\nCarD binds the non-template strand, which stabilizes the transcription bubble and\nfacilitates pause escape. This conclusion is supported by in vitro transcription assays\nshowing that CarD decreases the intensity of the +7 nt paused RNA band (Extended\nData Fig. 1H, I). We have revised the Results section (Page 17, Lines 341–347) to\nincorporate these findings and clarify the role of CarD.\n12. page 6 lines 101-104: authors refer to the studies on E. coli sig70-RNAP reporting pausing at +7 register. This is not\nrelevant because they use a different RNAP and a structurally distinct sigma factor.\nWe thank the reviewer for this point. We have performed new in vitro transcription\nexperiments using Mtb σE-RNAP, which confirm that initial transcription indeed pauses\nat the 6 to 7 register (Extended Data Fig. 1H, I). Although E. coli σ70-RNAP also\npauses at this position, our results demonstrate that this pause is conserved in Mtb\ndespite structural differences between the RNAPs and sigma factors. This supports\nthe relevance of our structural analyses of the paused complexes.\n13. page 9 lines 159 – 160: I see no evidence showing that CarD stabilized the non-template ssDNA of the transcription\nbubble.\nWe thank the reviewer for this thoughtful comment. In the paused transcription\ncomplex, we observed that CarD interacts with the non-template strand to stabilize the\ntranscription bubble. Using a fluorescence-labeled transcription bubble assay (Cordes\net al., Biochemistry 2010, 49: 9171-9180), we observed that CarD promotes\ntranscription bubble formation with σE, indicating stabilization of the RPo (Fig. 4G).\nConsistently, in vitro transcription assays show that CarD decreases the intensity of\nthe +7 nt paused RNA band, supporting its role in stabilizing the bubble and facilitating\npause escape (Extended Data Fig. 1H, I). Together with our structural observations\nof CarD binding to the non-template strand, these data support its role in transcription\nbubble stabilization and pause modulation.\n14. page 9 lines 163-165: authors wrote “It has been known for decades that RNAP adopts a pausing state when the\nnascent RNA extends to 6 to 7 nucleotides.“ If authors refer to the initial transcription pausing by E. coli sig70-RNAP, it has been published in\n2016. It’s not “for decades”.\nWe thank the reviewer for pointing this out, and we apologize that the wording in\nour previous manuscript was not rigorous. We agree that describing this as “for\ndecades” was an overstatement. The initial transcription pausing by E. coli σ70-RNAP\nwas first reported in 2016 (Duchi et al., Mol Cell 2016, 63: 939-950). We have revised\nthe text accordingly to state that this phenomenon was first described in 2016 (Page\n11, Lines 218-220).\n15. page 11 lines 196-197: authors wrote “… the σ3.2 domain of σE stimulates pausing through its insert into RNA exit\nchannel, forming a roadblock that promotes RNAP swiveling and interrupts nucleotide addition.”\nAs expalined above, there is no biochemical data supporting this conclusion.\nWe thank the reviewer for this important comment and apologize that our initial\nmanuscript did not include biochemical evidence to support this conclusion. We\nobserved that the σ3.2 region of σE inserts into the RNA exit channel and forms a steric\nroadblock. As the RNA extends, the 5′ end of the nascent transcript clashes with the\nσ3.2 region, and this steric interference promotes swiveling of RNAP, ultimately leading\nto transcriptional pausing. To validate this mechanism, we have now performed in vitro\ntranscription assays (Extended Data Fig. 1H, I), which confirm that Mtb σE-RNAP\npauses at the 7-nt stage. These biochemical data, together with our structural\nobservations, provide support for the revised description of σ3.2-mediated pausing.\nWe have updated the manuscript accordingly (Page 13, Lines 251-255).\n16. page 13 lines 241 – 242: authors wrote “The FRET assay validated this conformational change, consistent with\npreviously reported changes in E. coli. ” The ensemble FRET measurements performed by authors do not include essential\ncontrols which allows to validate it.\nWe appreciate the reviewer’s comment, and we apologize that the wording in our\nprevious manuscript was not rigorous. In the revised manuscript, we have clarified that\na DNA-only control was included in our ensemble FRET measurements, allowing us\nto exclude the possibility that the observed signal originated from free DNA rather than\nRNAP-bound complexes (Extended Data Fig. 7F). While we acknowledge that\nensemble FRET has inherent limitations compared with single-molecule approaches,\nour results are consistent with previously reported conformational changes in E. coli\nRNAP. We have revised the relevant text to clarify the controls and the scope of\ninterpretation (Page 15, Lines 293-298).\n17. page 13 lines 253 – 255: authors wrote “This minimal interface suggests a transient association mechanism at the\ntranscriptional checkpoint, distinct from stable binding.”\nIts not clear what do you mean under “transcriptional checkpoint” here ?\nWe thank the reviewer for raising this point. In our manuscript, by “transcriptional\ncheckpoint” we specifically refer to the decision point during initiation when RNAP\ncan either pause or proceed into productive elongation. At this stage, σE engages with\nRNAP through a minimal and transient interface, which differs from the more stable\nassociations observed in elongation complexes. To avoid ambiguity, we have revised\nthe text to clarify this meaning (Page 16, Lines 309-312). The revised sentence now\nreads: “This minimal interface suggests a transient association mechanism at the\ntranscriptional checkpoint, i.e., the stage where RNAP either pauses or escapes into\nproductive elongation, distinct from stable binding.”\n18. Extended data Figure 1j. The EMSA performed with the bubble template is not informative and adds nothing. It shows\nthat there is binding, but there is nothing to prouve that there is a complex with sigE-CarD-RNAP. RNAP core alone would\nbind and shift this template perfectly well. Also, I do not understand why so huge amounts of RNAP (18 mg) were used ?\nAlso, final concentration of proteins is not mentioned in the Methods section, while it should be.\nWe thank the Reviewer for critical questions and insightful suggestions. In the\nrevised manuscript, we have repeated the EMSA experiments with appropriate\ncontrols (σE, CarD, RNAP core, and σE-RNAP-CarD) to better illustrate the\ncontributions of each component. These new data are now included in Extended Data\nFigure 1G. we apologize for the mistake in reporting the amount of RNAP, which was\nincorrectly listed as 18 mg; the correct amount is 18 μg. In addition, we have updated\nthe Methods section to include the final concentrations of proteins used in the EMSA\nassays for clarity and reproducibility.\n19. Extended data Figure 1h . Is it expected that CarD will form a stable complex with sigE (peak2) ? How can it contribute to\nregulation ?\nWe appreciate the reviewer’s careful observation. Peak 2 does not represent a\nstable CarD-σE binary complex. Rather, it arises from the co-elution of excess CarD\nand σE added during assembly, which were not completely resolved during size-\nexclusion chromatography. We have clarified this point in the revised legend of\nExtended Data Fig. 1E to avoid confusion (Page 43, Lines 775-776).\nAlthough CarD does not form a stable complex with σE alone, it contributes to\nregulation by stabilizing the σE-RNAP-DNA complex. As shown by our FRET\nexperiments, CarD promotes transcription bubble stability (Fig. 4G), and our in vitro\ntranscription assays further demonstrate that CarD reduces the accumulation of the\n+7 paused RNA, facilitating pause escape (Extended Data Fig. 1H, I). These results\nsuggest that CarD functions as a transient co-factor: rather than binding stably to σE in\nisolation, it interacts with the transcription complex in a dynamic manner to modulate\npausing and initiation efficiency.\n20. Figure 3. There is a mismatch between Fig.3 panel lettering (panels g, h) and Fig.\n3 legend (panels j,k).\nWe thank the reviewer for pointing this out and apologize for the oversight in the\npanel labeling. The panel lettering and corresponding figure legend in Figure 3 have\nnow been corrected so that they match accurately.\n21. EMSA in Figure 3g is not informative. Which DNA was used ? Was there a\ncompetitor added ? What is a rational in doing EMSA with sigE alone ?\nWe thank the reviewer for this helpful comment. In the original EMSA, we used a\npre-melted DNA scaffold (Extended Data Fig. 1D) that was identical to the one\nemployed for assembling cryo-EM complexes, in order to maintain consistency\nbetween the biochemical and structural experiments. No competitor DNA was included\nat that stage, since our purpose was to test the direct binding capacity of σE and its\nmutant under simplified conditions. We also examined σE alone because when RNA\npolymerase is present, σE is largely embedded within the polymerase structure, and\nsimple point mutations may not produce detectable differences in DNA binding\ncompared to wild-type. Measuring σE on its own allowed us to assess the intrinsic DNA-\nbinding properties of the wild-type and mutant proteins\nTo address the reviewer’s concerns, we have now redesigned the EMSA\nexperiments. In the revised assays, we used fully complementary DNA (Extended\nData Fig. 1H) containing the canonical −10 motif (c/gGTTg/c) and compared wild-type\nand mutant σE both alone and in complex with RNAP. This new setup enables us to\nassess intrinsic DNA-binding while also evaluating the effect of RNAP on complex\nformation. The revised experiments, now included in Fig. 3G and Extended Data Fig.\n8E, provide clearer and more informative results that complement the structural\nanalyses.\n22. Figure 4g. A control without CarD is missing. I do not see how this experiment\nreveals CarD cooperativity .\nWe thank the Reviewer for critical questions and insightful suggestions. In the\nrevised experiments, we included a control without CarD to establish the baseline\ntranscription behavior. Compared with this control, addition of wild-type CarD\nsignificantly enhances transcription activity, and the CarD mutant also increases\nactivity but to a lesser extent, indicating partial retention of function (Fig. 4H). These\nresults demonstrate the cooperative effect of CarD in σE-dependent transcription.\nFurthermore, in vitro transcription pausing assays (Extended Data Fig. 1H, I) and\nFRET-based bubble stability measurements (Fig. 4G) both show that CarD cooperates\nwith σE to stabilize the transcription complex and facilitate escape from the paused\nstate. Together, these data provide multiple lines of evidence that CarD acts\ncooperatively with σE to modulate transcription, supporting our conclusions.\n23. Overall, manuscript should be rewritten in more concise from. Some sections from\nResults may go to Introduction (e.g. p12 lines 222- 227, p14, lines 261-265). Discussion is too excessive. Speculations on NusG and transcription -translation\ncoupling are irrelevant to the current study.\nWe thank the Reviewer for critical questions and insightful suggestions. We have\nmoved the sections highlighted (p12, lines 222-227; p14, lines 261-265) into the\nIntroduction (Page 4, Lines 59-65; Page 5, Lines 85-89). We have also consolidated\nsimilar content in the Discussion and streamlined the text to make the manuscript more\nconcise. The sections on NusG and transcription-translation coupling have been\nremoved, as they are not directly relevant to the present study. These revisions make\nthe manuscript more concise while retaining important mechanistic discussions\nrelevant to transcription initiation, CarD function, and σE retention.\nPoint-by-point responses\nWe sincerely thank the reviewers and editor for their thoughtful and constructive\ncomments, which have helped us substantially improve the manuscript. In this revised\nversion, we have carefully addressed all remaining issues through additional\nexperiments, clearer descriptions, and refined interpretations.\nIn this revision, we have:\n Performed new in vitro transcription assays with promoter scaffolds identical to\nthose used for cryo-EM, which confirmed a transcriptional pause at the +6/+7\nRNA register.\n Expanded the FRET analyses by providing detailed efficiency calculations and\ncorrelating the results with predicted structural transitions.\n Conducted a fluorescence-based assay demonstrating CarD-mediated\nstabilization of the σᴱ-RPo complex.\n Clarified figure legends, improved terminology and labeling, and removed\nspeculative statements to enhance clarity and precision.\n Revised the Results and Discussion sections to emphasize the transient and\nregulatory nature of pausing and the modulatory role of CarD.\nReferee's remarks are shown in black and our responses are in blue. All\nmodifications in the revised manuscript are highlighted in red.\nREVIEWER COMMENTS\nReviewer #1 (Remarks to the Author):\nThe authors have satisfactorily addressed all my previous concerns in the revised\nmanuscript. I have no further comments and fully support the publication of this study.\nWe thank the Reviewer for the generous and encouraging comments. We are\ndelighted that our revisions have addressed the reviewer’s concerns and that our work\nis now fully supported for publication.\nReviewer #2 (Remarks to the Author):\nThe authors have made substantial efforts to clarify many points in their response and\nthe revised manuscript, and addressed most issues raised in the reviews. The\nmanuscript is clearly interesting and making an important contribution. However, some\nkey questions and revisions remain:\nRe my main comment #1: the authors need to caveat their observation with the fact\nthat scrunching in their complexes is clearly very different from complexes prepared\nthrough an actual reaction. The fact that other works had used this strategy does not\ntake away the fact that they had the same caveat; I understand the necessity to get\nstable states, but the states may be different from the ones formed during the actual\nreaction, and this needs to be explicitly stated in the ms.\nWe thank the reviewer for this insightful comment emphasizing the distinction\nbetween our scrunched complexes and those formed during actual transcription\nreactions. We agree that our structures represent stabilized intermediates, which may\ndiffer in subtle ways from the transient states formed under native reaction conditions.\nWe have now explicitly stated this limitation in the Results section (page 15, lines\n291–300) and further discussed it in the Discussion section (page 20, lines 388–398),\nemphasizing that although such stabilized complexes have been widely used in\nprevious structural studies, they may not fully recapitulate the dynamic nature of\nscrunching during active transcription.\nRe my main comment #5. The FRET assay remains poorly described. What is the\nFRET efficiency of the various samples? What is the prediction from the structure?\nWhy is the FRET efficiency different only for RNA7? And not for RNA6 or RNA8? This\nassay needs a proper introduction, predictions, description, and discussion, as well as\na better illustration.\nWe thank the reviewer for this insightful comment and apologize for the insufficient\ndescription in the previous submission. In the current revision, we have defined FRET\nefficiency as the normalized change in Cy3 fluorescence relative to the DNA-only\ncontrol. The detailed calculation procedure is now described in the Methods section\n(page 30, lines 602–607) and added corresponding explanations in the legend of\nExtended Data Fig 7F (page 56, lines 910–919). In the Results section (page 16,\nlines 305–315), we further describe how the measured FRET efficiencies correspond\nto the structural transitions observed during RNA extension. Specifically, structural\nanalysis suggests that elongation of the RNA induces a steric clash with the σ3.2\nregion, resulting in conformational rearrangements that elevate the upstream DNA and\nthus reduce FRET efficiency. At the 7-nt RNA stage, the complex exists as a mixture\nof paused and non-paused states, resulting in a relatively lower FRET efficiency. In\ncontrast, RNA5 and RNA6 have not yet reached the σ3.2 region and therefore show\nrelatively higher FRET efficiencies, while RNA8 and RNA9 likely overcome the σ3.2\nbarrier or undergo backtracking, resulting in increased FRET again. These\ninterpretations are consistent with our structural observations, and we have clarified\nthem in the Results section (page 16, lines 305–315). In addition, the corresponding\nFRET efficiency data have been added to Extended Data Fig. 7F to illustrate these\ntrends.\nWe thank the reviewer for this insightful comment and apologize for the insufficient\ndescription and lack of clarity in the previous submission. To address the reviewer’s\nsuggestion for a clearer introduction, predictions, description, and discussion of the\nFRET assay, we have comprehensively revised the relevant text throughout the\nmanuscript. Specifically, we have introduced the assay rationale and FRET efficiency\ncalculation in the Methods section (page 30, lines 602–607). We have also included\nstructure-based predictions of FRET efficiency changes and described the\ncorrespondence between structural rearrangements and fluorescence signals in the\nResults section (page 16, lines 305–315). Furthermore, we have discussed the\nlimitations of the current bulk FRET measurements and the need for further validation\nin the Discussion (page 29, lines 498–504). In addition, we have added a schematic\nillustration and explanatory notes in Extended Data Fig. 7 to clarify the FRET\nmechanism, summarize the experimental design, and visualize the relationship\nbetween structural rearrangements and FRET efficiency changes.\nRe my minor comment #7: Fig 1I needs to be annotated so we can understand what\nthe “SigE” lane is. Further, if the sigE lane is for the run-off reaction, then the band that\naccumulates is the 6-mer, not the 7-mer (as in Duchi et al Mol Cell 2016 for the E.coli\nsystem), hence the pause (i.e., the point in synthesis where the conversion from n to\nn+1 occurs) occurs at the 6-mer level. As a result, the discussion and model need to\nadapt to accommodate the functional data.\nWe thank the reviewer for the helpful comment and apologize for the unclear\nlabeling in the previous submission. In the current revision, we have now corrected the\nannotation in Extended Data Fig. 1I and clarified in the figure legend that the “SigE”\nlane represents the run-off transcription reaction.\nWe thank the reviewer for this careful observation and apologize that our previous\nrevision did not sufficiently integrate the structural and biochemical views of pausing\nat the 6-nt to 7-nt stage. Transcriptional pausing during initial transcription is inherently\ndynamic, and complexes are expected to sample multiple, closely related\nintermediates rather than a single discrete state. In our cryo-EM analysis, the paused\nconformation that we capture at a nominal RNA length of 7-nt adopts a half-\ntranslocated architecture: the RNA has reached 7-nt, whereas the template DNA in the\nactive site remains in a pre-translocated register equivalent to the 6-nt position (Fig.\n1F). This half-translocated arrangement is a well-recognized hallmark of paused RNAP\ncomplexes. Therefore, we propose that pausing occurs during the transition between\nthe 6-nt and 7-nt RNA lengths, rather than being confined to a specific RNA length of\neither 6-nt or 7-nt.\nConsistent with this, our in vitro transcription assays show two species at 6-nt and\n7-nt, with the 6-nt band more prominent and the 7-nt band weaker. Taken together, the\nstructural and biochemical data suggest that pausing occurs during the transition from\n6-nt to 7-nt RNA synthesis, rather than being strictly confined to a single RNA length.\nIn the revised manuscript, we therefore do not assign the pause to “6-nt” or “7-nt” alone,\nbut instead describe it as a checkpoint spanning the 6-nt to 7-nt transition. This\ninterpretation is in line with the kinetic analysis by Duchi et al. (Mol Cell 2016, 63: 939–\n950), where extension from 6-nt to 7-nt was found to be rate-limiting and elongation\nbeyond 7-nt proceeded more rapidly. We have revised the relevant text in the Results\n(page 13, lines 251–253) and Discussion (page 24, lines 476–479) to reflect this view\nof a dynamic 6-nt to 7-nt checkpoint rather than a pause assigned to a single register.\nI also think that the term “allosteric” is still a bit confusing, and best to avoid. The term\n“RNA-induced” seems both adequate and appropriate.\nWe sincerely thank the reviewer for the helpful suggestion and apologize for the\ninaccurate terminology used in our previous version. The term “allosteric” did not\nprecisely capture the mechanistic nature of the process we describe. We appreciate\nthe reviewer’s clarification and fully agree that “RNA-induced” provides a more\naccurate and appropriate description of the conformational changes observed.\nAccordingly, we have revised the manuscript to replace “allosteric” with “RNA-induced”\nthroughout the relevant sections to ensure terminological accuracy and conceptual\nclarity.\nReviewer #3 (Remarks to the Author):\nWhile the revised manuscript has been improved my principal concerns have not been\nresolved.\n1. My point was that the authors provided no evidence for pausing at the +6/+7\nregisters on the promoter templates used for cryo-EM. In the revised version, the\nauthors have added a run-off transcription assay (Extended Data Fig. 1H,I). However,\nupon reading, it appears that this assay was performed using a different template and\nunder different conditions: including a different -10 sequence, dsDNA bubble instead\nof a mismatch bubble, and de novo initiation instead of RNA primers. Since pausing is\nhighly dependent on the DNA template architecture, I do not see how this experiment\ncan support the structures presented by the authors. In essence, there are structures\nand there is a functional assay, but they are disconnected.\nFinally, a single run-off data point does not demonstrate pausing at +6/+7. Although\nthere are weak bands corresponding to 6-nt and 7-nt RNAs, these could simply\nrepresent abortive products. The only proper way to demonstrate pausing is to monitor\nthe kinetics of promoter escape: if the +6/+7 bands first increase and then disappear,\none could then conclude that a pause occurs.”\nFurthermore, in the Figure legend, authors should better explain what is shown on the\ngel. What is in the “control” line ? How lines with 6nt-pCp AF647 and 7nt-pCp AF647\nwhere obtained ? Are these just RNA primers alone or in complex with RNAP ?\nWe thank the reviewer for this insightful comment and for emphasizing the\nimportance of directly correlating biochemical assays with the structural analysis. In\nthe current revision, we have performed additional in vitro transcription assays using a\npre-melted promoter DNA scaffold identical to that used for cryo-EM sample\npreparation (Extended Data Fig. 1J, K). Because fixed RNA primers could bias the\ndetection of potential paused intermediates by predefining the transcript length, we\ninstead employed de novo transcription reactions to more accurately capture all RNA\nspecies generated during initiation. To maintain full consistency with the cryo-EM\nsamples, the DNA scaffolds used in these assays were relatively short, which resulted\nin run-off transcripts migrating close to the paused RNA species on the gel (Extended\nData Fig. 1 K). These new time-course experiments were designed to provide kinetic\nevidence of pausing at the +6/+7 registers, and the results indeed show a gradual\nreduction in the intensity of the 6-nt and 7-nt RNA bands as the reaction proceeds,\nparticularly between 30 and 60 minutes (Extended Data Fig. 1K). Because\ntranscription initiation pausing is inherently coupled with abortive release, the reaction\ninvolves overlapping processes that make it difficult to obtain a sharply defined kinetic\ntrend. We also tested shorter reaction intervals, but the RNA yield was too low and the\nbands became less discernible. Nevertheless, the overall pattern supports the\ninterpretation that a transient paused intermediate exists during early transcription\ninitiation. Moreover, σE+CarD reactions show reduced accumulation of paused\nproducts, consistent with the results obtained using the fully matched promoter\ntemplate (Extended Data Fig. 1I, K).\nIn the current revision, we have revised the figure legends (page 45, lines 813–\n833) to clarify the composition of the “control” lanes and the preparation of the AF647-\nlabeled 6-nt and 7-nt RNA markers. The “control” lanes represent reactions performed\nwithout promoter DNA, while all other components were identical to the σE+CarD\nreaction, allowing us to assess background RNA signals. The 6-nt-pCp–AF647 and 7-\nnt-pCp–AF647 markers were prepared by labeling synthetic RNA oligonucleotides of\n6 and 7 nucleotides in length with pCp-AF647, following the procedure described in\nExtended Data Fig. 1H. All RNA species analyzed in these assays represent RNA\noligonucleotides alone rather than RNAP-RNA complexes, as fluorescence labeling\nwas performed after TRIzol extraction of RNA products from the completed reactions\nprior to gel analysis (Extended Data Fig. 1H, J). Therefore, no RNA primers were\npresent in complex with RNAP, and the labeled RNA alone was analyzed on denaturing\nurea PAGE gels. Additionally, we have slightly adjusted the phrasing in the Results\nsection (page 18, lines 360–363) to state that the observed structural features are\n“consistent with the trends observed in our in vitro transcription assays,” reflecting the\ntransient and partially overlapping nature of pausing and abortive events.\n2. I their response letter authors explain the rational for using CarD by analogy with\nCarD from Myxococcus xanthus which has been shown to regulate ECF sigmas\ntranscription. While CarD is indeed a conserved protein, Myxococcus xanthus is not\nan Actinobacteria, as claimed by authors, and such comparison is inappropriate.\nSee:\nhttps://www.ncbi.nlm.nih.gov/Taxonomy/Browser/wwwtax.cgi?id=34\nWe thank the reviewer for this valuable correction and apologize for the inaccurate\ndescription in the previous submission. In the current revision, we have removed the\ntaxonomic reference and rephrased the statements to avoid any direct comparison\nbetween Myxococcus xanthus and Mycobacterium tuberculosis (page 5, lines 86–89).\nThe revised text now simply notes that CarD has been reported to modulate\ntranscription mediated by an ECF σ factor in M. xanthus (Ref. 46), and highlights that\nits potential role in σᴱ-dependent transcription in M. tuberculosis has not been\ncharacterized before. This change accurately reflects the published evidence and\navoids overinterpretation.\n3. Its not clear how CarD can stimulate pause escape if it stabilizes interaction with the\nssDNA of transcription bubble. In that case one would expect an opposite: CarD will\nstabilize pause.\nWe thank the reviewer for this important comment. Indeed, previous studies have\ndemonstrated that CarD stabilizes the open promoter complex (RPo) and prevents\nbubble collapse. These findings are consistent with our observation that CarD\nenhances σᴱ-dependent transcription initiation.\nOur results indicate that during the transition from 6-nt to 7-nt RNA synthesis,\nCarD reduces the lifetime of the paused state (Extended Data Fig. 1I, K). In our\nstructural analysis, we observe that CarD binds to the template strand in the initiation\ncomplex, while it binds to the non-template strand during the paused state (Fig. 4A-\nD). In both scenarios, CarD serves to stabilize the transcription bubble, although the\nbinding mode differs. Specifically, during initiation, CarD stabilizes the transcription\nbubble to facilitate the unwinding of the DNA duplex and prevent collapse. During the\npaused state, CarD stabilizes the scrunched transcription bubble, helping to ensure\nthe transition from the paused state to productive elongation. This dual role is\nconsistent with our biochemical experiments, which show that CarD reduces the\nintensity of the 6-nt and 7-nt paused bands (Extended Data Fig. 1H, I). Therefore, we\npropose that CarD’s function in transcription initiation is to stabilize the transcription\nbubble throughout, first enabling efficient DNA opening and later stabilizing the\nscrunched bubble to support the transition to elongation, rather than prolonging the\npause.\nWe have updated the Results (page 19, lines 360–372) and Discussion (page\n24, lines 480–485) sections to clarify this dual functionality of CarD, emphasizing its\nrole in both initiation and pausing, and to better explain how it stabilizes the\ntranscription bubble in a conformation conducive to transcription elongation.\n4. Fig. 4G FRET assay does not directly demonstrate any effect on bubble ‘stability’.\nIt only shows an increase in fluorescence, which the authors interpret as an effect on\nstability. If the authors wish to make a stronger claim regarding RPo stability, they\nshould perform a dissociation kinetics study, as I suggested during the first round of\nrevision. This analysis could also be carried out using FRET. As a more simple\nfluorescent assay they can use promoter with Cy3 probe in transcription bubble (e.g.\nDOI: 10.1093/nar/gkw577 )\nWe thank the reviewer for this constructive suggestion and for pointing us to the\nrelevant reference (DOI: 10.1093/nar/gkw577). Following this recommendation, we\nhave re-examined the effect of CarD on RPo formation using a fluorescence-based\nkinetic assay inspired by this method. Although a stopped-flow instrument was not\navailable to achieve millisecond resolution, we performed a time-resolved fluorescence\nassay with second-level temporal resolution (Fig. 4G). The results consistently show\nthat the presence of CarD leads to a higher plateau compared to the RNAP-σE-only\ncomplex, indicating that CarD stabilizes the σE-RPo complex and enhances its steady-\nstate accumulation. This new dataset has been included in the revised Fig. 4G and\ndescribed in the Methods (page 31, lines 617–625) and Results (page 19, lines 365–\n372) sections.\n5. p4 lanes 65-71: speculations about weak affinity of sigE to Mtb RNAP are\nunwarranted. You cannot, and should not, draw conclusions about the relative affinities\nof sigA versus ECF sigma factors for M. tuberculosis (Mtb) RNAP based on the\ncorresponding affinities of sig70 versus ECF sigma factors in E. coli. Numerous studies\nover the past decades have clearly shown that Mtb RNAP behaves differently from E.\ncoli RNAP; for example, sigA exhibits only weak affinity for Mtb RNAP in the absence\nof RbpA.\nWe thank the reviewer for raising this point and fully agree that inferring σE-RNAP\nbinding affinity in Mtb based on published data from E. coli was inappropriate. We\napologize for this speculative interpretation. Since this inference is not essential to our\nconclusions, we have removed the statement regarding relative binding affinity\nbetween σ factors (page 4, lines 63–67). The revised text now focuses only on the\nrationale for choosing σE — its simpler domain architecture and suitability for capturing\ninitiation intermediates — without making assumptions about RNAP-σ binding\nstrengths.\n6. p. 17 “line 343 “...CarD ... facilitates pausing at the 6-7 nt RNA stage”.\nIt contradicts to your pervious conclusion that CarD suppresses pausing.\nWe apologize for the confusion caused by the previous wording. We thank the\nreviewer for carefully noting this inconsistency. In our revised manuscript, we have\nclarified that CarD does not directly facilitate pausing but rather modulates the paused\nstate. Our structural analysis indicates that CarD binds to the template strand during\ninitiation, and to the non-template strand during pausing, playing a stabilizing role in\nboth cases. Specifically, in the initiation state, CarD prevents premature collapse of the\ntranscription bubble, whereas in the paused state, it stabilizes the scrunched\ntranscription bubble, facilitating the transition from pausing to elongation. This dual role\nis consistent with our biochemical data, which show that in the presence of CarD,\nthe 6-nt and 7-nt paused bands become weaker (Extended Data Fig. 1H, I), supporting\nthe idea that CarD reduces the duration of pausing and promotes escape from the\npaused state.\nIn the revised manuscript, we have made the necessary changes to the Results\n(page 19, lines 365–372) and Discussion (page 24, lines 480–485) sections to ensure\nour interpretation aligns with the structural and biochemical data. This correction now\nreflects that CarD's role is not to directly facilitate pausing, but rather to modulate the\npaused state and promote transition into productive elongation.\nPoint-by-point responses\nWe sincerely thank the reviewers and the editor for their thoughtful and constructive\ncomments, which have helped us further improve the clarity and rigor of the manuscript.\nIn this revised version, we have carefully addressed all remaining points raised by the\nreview through focused clarification and minor textual revisions.\nIn this revision, we have:\n• Removed the ensemble FRET dataset, as technical limitations made its interpretation\nuncertain and the data were not required for our mechanistic conclusions.\n• Clarified the correspondence between the structural and biochemical complexes,\nincluding how the matched +1 to +7 scaffold relates the structures to de novo initiation,\nand added a concise explanation of the inherent limitations of de novo assays at the\n6–7 nt stage.\n• Refined the presentation of the initiation-paused intermediate and the role of CarD\nthroughout the Abstract, Introduction, Results, and Discussion, ensuring that our\ninterpretations rely strictly on structural and biochemical evidence and avoiding\noverstating the contribution of any single factor.\nReferee's remarks are shown in black and our responses are in blue. All\nmodifications in the revised manuscript are highlighted in red.\nReviewer #2 (Remarks to the Author):\nThe revised manuscript is much improved, and several key conclusions are now more\nclearly supported.\nWe thank the reviewer for this encouraging comment and are pleased that the\nrevised manuscript more clearly supports our key conclusions.\nRegarding the transition from IC6 to IC7, the authors’ discussion is accurate, and\ntheir reference to associated literature is relevant and appropriate; the paused state is\nan intermediate that can then give rise to different paths; this path distribution may be\ndifferent in Mtb vs E.coli, but the gels qualitative support the presence of states that\ngive rise to 6-mer (and 7-mer) RNAs, likely due to abortive initiation. I also note that\nthere is very recent work (doi.org/10.1101/2025.11.27.690701) on this transition and\nthe paths emanating from it, as well as the sequence dependence of pausing, which\nthe authors may want to cite to further support their argument.\nWe thank the reviewer for the positive assessment that our discussion of the IC6 to\nIC7 transition is accurate and that the cited literature is relevant and appropriate. We\nare pleased that the reviewer agrees with our interpretation that the paused state\nrepresents a transient intermediate that can give rise to multiple downstream pathways,\nand that our gels qualitatively support the presence of states leading to 6-nt and 7-nt\nRNA species, likely reflecting abortive initiation from this checkpoint.\nWe thank the reviewer for pointing out this very recent preprint (doi:\n10.1101/2025.11.27.690701). We agree that this study provides additional support for\nthe view that the 6–7 nt transition represents a branching intermediate during initiation,\nwith outcomes influenced by sequence context and kinetic partitioning. We have now\ncited this work (Reference 65) in the Discussion (Page 22, Lines 431-433) and briefly\nnoted its consistency with our interpretation.\nThe removal of the FRET data is a positive change, as the previous analysis was\nconvoluted, had many caveats and distracted from the main conclusions.\nWe thank the reviewer for this positive assessment. We agree that removal of the\nensemble FRET data improves the clarity and focus of the manuscript, and we are\npleased that the reviewer feels this change helps to better highlight the main structural\nand mechanistic conclusions.\nThe interpretations related to the premelted and fully double-stranded DNA substrates\nis also robust and strengthens the overall model.\nWe thank the reviewer for this encouraging assessment. We are pleased that the\ninterpretation of the premelted and fully double-stranded DNA substrates is considered\nrobust and that it is viewed as strengthening the overall mechanistic model.\nThe CarD effect (EDFig. 1I) itself is clear and well supported. One point that warrants\nbrief clarification is the high run-off to pause ratio (EDFig. 1I). The long reaction time\n(30 min) may contribute to this effect, and a short discussion of possible causes would\nbe helpful. The authors may also wish to comment on the identity of the major band\nbetween the run-off product and the 6–7 nt species.\nWe thank the reviewer for recognizing that the CarD effect is clear and well\nsupported. We agree that the relatively high run-off to pause ratio is likely influenced\nby the long reaction time (30 min), which was chosen to allow sufficient accumulation\nof the transient 6–7 nt paused species for reliable detection. We now briefly note in the\nDiscussion (Page 23, Lines 455-458) that the relatively long incubation time allows\ncontinued transcript extension and accumulation of longer RNA products, which can\nlead to an increased proportion of run-off RNA relative to short paused species, without\naltering the identification or stability of the 6–7 nt initiation-paused intermediate.\nRegarding the prominent band between the run-off product and the 6–7 nt species,\nwe believe this most likely corresponds to a short, prematurely terminated or transiently\nstalled elongation product formed during the early stages of transcription initiation.\nImportantly, its presence does not affect the identification of the 6–7 nt paused species\nor the interpretation of the CarD-dependent stabilization of the initiation-paused\nintermediate. We have added a short comment in the Discussion (Page 24, Lines 466-\n469) to acknowledge this band and to clarify that it does not alter our mechanistic\nconclusions."}]

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