Escherichia coli RNAP, with or without TraR, and of an RNAP-promoter complex. TraR binding induced RNAP conformational changes not seen in previous crystallographic analyses, and a quantitative analysis revealed TraR-induced changes in RNAP conformational heterogeneity. These changes involve mobile regions of RNAP affecting promoter DNA interactions, including the βlobe, the clamp, the bridge helix, and several lineage-specific insertions. Using mutational approaches, we show that these structural changes, as well as effects on σ70 region 1.1, are critical for transcription activation or inhibition, depending on the kinetic features of regulated promoters.TraR and its homolog DksA are bacterial proteins that regulate transcription initiation by binding directly to RNA polymerase (RNAP) rather than to promoter DNA. Effects of TraR mimic the combined effects of DksA and its cofactor ppGpp, but the structural basis for regulation by these factors remains unclear. Here, we use cryo-electron microscopy to determine structures of  Cells need to make proteins in order to survive. To make a protein, a cell first needs to take the information from a gene coded in its DNA and copy it into a template made of a different molecule called RNA via a process called transcription. Not all genes are transcribed or expressed at the same time or the same rate, because a cell needs different proteins depending on its environment. The molecules that regulate when each gene is transcribed are called transcription factors.Escherichia coli is starved for the building blocks it needs to make proteins, it changes the expression of almost one quarter of its genes within 5 minutes. This broad response requires two transcription factors, ppGpp and DksA. Unlike most transcription factors, these two molecules bind directly to the enzyme responsible for transcribing DNA into RNA, the RNA polymerase, but they do not bind to DNA.When the bacterium Another transcription factor called TraR can mimic the combined effects of ppGpp and DksA on transcription, changing the conformation of RNA polymerase in the same way. Now, Chen et al. have used a high-resolution imaging technique called cryo-electron microscopy to reveal the details of how the structure of RNA polymerase changes in response to TraR binding, and by analogy, in response to ppGpp and DksA.The experiments showed that TraR interacts with regions of RNA polymerase that move when the protein binds DNA. These interactions can either help or hinder the start of transcription depending on which DNA sequence the RNA polymerase binds. Using the structures obtained via cryo-electron microscopy as guides, Chen et al. mutated each of the mobile parts of RNA polymerase individually to determine which interactions with TraR were necessary to change gene expression.These results shed light on a fundamental process in all living cells, the initiation of transcription, and how it changes in response to the cell's nutritional environment. This may help explain how different cells regulate gene expression and may also lead to the development of new antibiotics. Escherichia coli (Eco), the essential primary σ factor, σ70, binds to RNAP to form the σ70-holoenzyme (Eσ70) that is capable of recognizing and initiating at promoters for most genes. Upon locating the promoter, Eσ70 melts a ~ 13 bp segment of DNA to form the open promoter complex (RPo) in which the DNA template strand (t-strand) is loaded into the RNAP active site, exposing the transcription start site  performs all transcription. In art site . A key fart site .Eco to reprogram bacterial metabolism in response to nutritional stresses during the so-called stringent response. Following amino acid starvation, ppGpp is synthesized by the RelA factor in response to uncharged tRNAs in the ribosomal A site  and ribosomal protein promoters , and activates amino acid biosynthesis and transport promoters  in vivo and in vitro , and Eσ70 bound to a promoter inhibited by TraR  contained approximately 41% of the particles and resolved to a nominal resolution of 3.7 Å (70(II)] contained approximately 33% of the particles and resolved to a nominal resolution of 3.8 Å (70(III)] contained the remaining 26% of the particles and resolved to a nominal resolution of 3.9 Å , interacting with key RNAP structural elements surrounding the active site, including the -NADFDGD- motif that chelates the active site Mg2+ (N (TraR residue S2) is only 4.3 Å from the active site Mg2+ . Indeed, the cryo-EM and X-ray structures superimpose with an rmsd of 4.26 Å over 3,471 α-carbons, indicating significant conformational differences.The interactions of TraR70 resolved to a nominal resolution of 4.1 Å  fold (70(I) and TraR-Eσ70(II), Si3 is in two distinct positions with respect to the RNAP (70(III) Si3 is disordered (70(I) structure [Si3(I)] interacts primarily with the β’shelf (SBHMa) and the β’jaw (SBHMb) in a manner seen in many previous Eco RNAP X-ray (70(II) structure [Si3(II)] is rotated 121° such that SBHMa interacts with the β’jaw and SBHMb interacts with TraRG (The three TraR-EσHM) fold , SBHMa aHM) fold . Si3 is the RNAP  and 3A, sordered . Si3 in AP X-ray  and cryoAP X-ray . Si3 in th TraRG , a disporrnB P1 and rpsT P2 ] is relevant to TraR function, we compared TraR-mediated function with wild-type (WT) and ΔSi3-RNAPs at promoters where TraR inhibits or activates transcription. Deletion of Si3 had little to no effect on TraR-mediated inhibition of  rpsT P2  but tranG residues  are central to the Si3-TraRG interface . Maximal inhibition was achieved at higher E46A or R49A TraR concentrations. However, these same variants exhibited at least ~2 fold reduced activation at the thrABC promoter (Three TraRnterface . Individpromoter  even at promoter . By contpromoter .G interaction enabled by the Si3(II) conformation accounts for part of the TraR-mediated effect on activation.The combination of the TraR-Si3 interface and the ΔSi3-RNAP mutants was epistatic; the same ~2 fold reduction in activation was observed as with the Si3-TraR interface mutants or the ΔSi3-RNAP mutant individually . These r70 without nucleic acids, this channel is occupied by the σ701.1 domain, which is ejected upon entry of the downstream duplex DNA  and an activated (thrABC) promoter with holoenzyme lacking σ701.1 (EΔ1.1σ70).The rotation of the βlobe-Si1 widens the gap between the βprotrusion and the βlobe  and chan70 exhibited weak transcription from the thrABC promoter in the absence of TraR , and transcription from this promoter was stimulated about ~4 fold in the presence of TraR  compared to WT-Eσ70 activity . We propose that TraR-mediated stimulation of σ701.1 release still occurs at inhibited promoters like rrnB P1 and rpsT P2, but this has little effect on transcription because these promoters are limited by their unstable RPo , so a random subset of particles from the other datasets (TraR-Eσ70 and Eσ70) were processed so that each map for multi-body refinement was generated from the same number of particles . The final maps used for multi-body refinement had nominal resolutions of 4.0 Å  of clamp changes were revealed  were represented in major components of clamp changes for the TraR-Eσ motions . As expe motions . See DisEco RNAP transcription initiation by binding to and altering the discrete conformation, as well as the conformational heterogeneity, of the RNAP in four major ways: (1) manipulation of the disposition of β'Si3  matches the Eσ70 crystal structure . Peak fractions were combined, treated with ULP1 SUMO-protease overnight, and dialyzed against 20 mM Tris-HCl, pH 8.0, 5% glycerol (v/v), 0.1 mM ethylenediaminetetraacetic acid (EDTA), 500 mM NaCl, 10 µM ZnCl2, 2.5 mM DTT, resulting in a final imidazole concentration of 25 mM. The ULP1-cleaved sample was loaded onto one 5 mL HiTrap IMAC HP column to remove His10-SUMO-tag along with any remaining uncut TraR. Tagless TraR was collected in the flowthrough and concentrated by centrifugal filtration . The sample was purified in a final step on a HiLoad 16/60 Superdex 200 column . Purified TraR was concentrated to 16 mg/mL by centrifugal filtration, flash-frozen in liquid N2, and stored at −80°C.The His10  was co-transformed with a pACYCDuet-1 plasmid containing Eco rpoZ into Eco BL21(DE3). The cells were grown in the presence of 100 µg/mL ampicillin and 34 μg/mL chloramphenicol to an OD600 of 0.6 in a 37°C shaker. Protein expression was induced with 1 mM IPTG  for 4 hr at 30°C. Cells were harvested by centrifugation and resuspended in 50 mM Tris-HCl, pH 8.0, 5% glycerol (v/v), 10 mM DTT, 1 mM PMSF, and 1x PIC. After French Press lysis at 4°C, the lysate was centrifuged twice for 30 min each. Polyethyleneimine  was slowly added to the supernatant to a final concentration of ~0.6% PEI with continuous stirring. The mixture was stirred at 4°C for an addition 25 min, then centrifuged for 1.5 hr at 4°C. The pellets were washed three times with 50 mM Tris-HCl, pH 8.0, 500 mM NaCl, 10 mM DTT, 5% glycerol (v/v), 1 mM PMSF, 1x PIC. For each wash, the pellets were homogenized then centrifuged again. RNAP was eluted by washing the pellets three times with 50 mM Tris-HCl, pH 8.0, 1 M NaCl, 10 mM DTT, 5% glycerol (v/v), 1x PIC, 1 mM PMSF. The PEI elutions were combined and precipitated with ammonium sulfate overnight. The mixture was centrifuged and the pellets were resuspended in 20 mM Tris-HCl, pH 8.0, 1 M NaCl, 5% glycerol (v/v), 5 mM DTT. The mixture was loaded onto three 5 mL HiTrap IMAC HP columns for a total CV of 15 ml. RNAP(β'-PPX-His10) was eluted at 250 mM imidazole in Ni-column buffer. The eluted RNAP fractions were combined and dialyzed against 20 mM Tris-HCl, pH 8.0, 100 mM NaCl, 5% glycerol (v/v), 5 mM DTT. The sample was then loaded onto a 35 mL Biorex-70 column , washed with 10 mM Tris-HCl, pH 8.0, 0.1 mM EDTA, 5% glycerol (v/v), 5 mM DTT] in a gradient from 0.2 M to 0.7 M NaCl. The eluted fractions were combined, concentrated by centrifugal filtration, then loaded onto a 320 mL HiLoad 26/600 Superdex 200 column  equilibrated in gel filtration buffer . The eluted RNAP was supplemented with glycerol to 20% (v/v), flash frozen in liquid N2, and stored at −80°C.A pET-based plasmid overexpressing each subunit of RNAP  as well as β'-PPX-HisEco His10-SUMO-σ70 was transformed into Eco BL21(DE3) by heat shock. The cells were grown in the presence of 50 µg/mL kanamycin to an OD600 of 0.6 at 37°C. Protein expression was induced with 1 mM IPTG for 1 hr at 30°C. Cells were harvested by centrifugation and resuspended in 20 mM Tris-HCl, pH 8.0, 5% glycerol (v/v), 500 mM NaCl, 0.1 mM EDTA, 5 mM imidazole, 0.5 mM 2-mercaptoethanol (BME), 1 mM PMSF, 1x PIC. After French Press lysis at 4°C, cell debris was removed by centrifugation. The lysate was loaded onto two 5 mL HiTrap IMAC HP for a total CV of 10 ml. His10-SUMO-σ70 was eluted at 250 mM imidazole in 20 mM Tris-HCl, pH 8.0, 500 mM NaCl, 0.1 mM EDTA, 5% glycerol (v/v), 0.5 mM BME. Peak fractions were combined, cleaved with ULP1, and dialyzed against 20 mM Tris-HCl pH 8.0, 500 mM NaCl, 0.1 mM EDTA, 5% glycerol (v/v), 0.5 mM BME, resulting in a final imidazole concentration of 25 mM. The cleaved sample was loaded onto one 5 mL HiTrap IMAC HP to remove His10-SUMO-tag along with any remaining uncut σ70. Tagless σ70 was collected in the flowthrough and concentrated by centrifugal filtration. The sample was then loaded onto a HiLoad 16/60 Superdex 200 in gel filtration buffer. Peak fractions of σ70 were pooled, supplemented with glycerol to a final concentration of 20% (v/v), flash-frozen in liquid N2, and stored at −80°C.Plasmid encoding 70 was formed by mixing purified RNAP and 2.5-fold molar excess of σ70 and incubating for 20 min at 37°C. Eσ70 was purified on a Superose 6 Increase 10/300 GL column  in gel filtration buffer . The eluted Eσ70 was concentrated to ~10 mg/mL (~21 μM) by centrifugal filtration (Amicon Ultra).Eσ70 was formed by mixing purified RNAP and a 2-fold molar excess of σ70 and incubating for 15 min at room temperature. Eσ70 was purified over a Superose 6 Increase 10/300 GL column in gel filtration buffer. The eluted Eσ70 was concentrated to ~5.0 mg/mL (~10 μM) by centrifugal filtration. Purified TraR was added (5-fold molar excess over RNAP) and the sample was incubated for 15 min at room temperature. An rrnB P1 promoter fragment  was added (2-fold molar excess over RNAP) and the sample was incubated for a further 15 min at room temperature. The rrnB P1 promoter fragment did not bind to TraR-Eσ70 under the cryo-EM grid preparation conditions - the subsequent structural analyses did not reveal any evidence of promoter binding.Eσ70 was prepared as described for TraR-Eσ70, but after the size exclusion chromatography the complex was concentrated to ~10 mg/mL (~20 μM) by centrifugal filtration. Duplex rpsT P2 promoter fragment  were glow-discharged for 20 s prior to the application of 3.5 μL of the samples. Using a Vitrobot Mark IV , grids were blotted and plunge-froze into liquid ethane with 100% chamber humidity at 22°C.CHAPSO {3-([3-cholamidopropyl]dimethylammonio)−2-hydroxy-1-propanesulfonate}  was added to the samples to a final concentration of 8 mM . The finhttp://www.mrc-lmb.cam.ac.uk/kzhang/) using a 2D template. Picked particles were extracted from the dose-weighted images with RELION  using a cryoSPARC ab-initio reconstruction (70(I) and TraR-Eσ70(II) were used for RELION multi-body refinements to examine clamp motions (Grids were imaged using a 300 keV Krios (FEI) equipped with a K2 Summit direct electron detector . Datasets were recorded with Serial EM  with a ph RELION  using a truction  generatetruction . The pol motions . Local r motions .70 image acquisition and processing were the same as for TraR-Eσ70 except with the following differences. Grids were imaged using a 200 keV Talos Arctica (FEI) equipped with a K2 Summit direct electron detector. Datasets were recorded with a pixel size of 1.3 Å over a defocus range of −1.0 μm to −2.5 μm. Movies were recorded in counting mode at 8.4 electrons/physical pixel/second in dose-fractionation mode with subframes of 0.2 s over a 10 s exposure (50 frames) to give a total dose of 84 electrons/physical pixel. Picked particles were extracted from the dose-weighted images in RELION  resulting in a subset of 479,601 particles for the first group and 329,293 particles for the second group. Curated particles were combined and a consensus refinement was performed in RELION using the cryoSPARC generated initial model resulting in a map with nominal resolution of 4.54 Å (without post-processing). Particles from this refinement  to give a total dose of 80 electrons/physical pixel. The rpsT P2-RPo dataset consisted of 6912 motion-corrected images with 973,481 particles. In RELION, a consensus refinement was performed using the extracted particles and a cryoSPARC generated initial model resulting in a 4.62 Å resolution map. Using the refinement parameters, 3D classification (N = 2) was performed on the particles without alignment. Classification revealed a lower resolution class and a higher resolution class, the latter with 370,965 particles with nominal resolution of 4.38 Å after RELION 3D auto-refinement. Refinement metadata and post-processing were used as inputs for RELION CTF refinement and RELION Bayesian Polishing  host cells. TraR and variants were purified as described  was used to induce expression of TraR (WT or variant) from escribed , either eviously . The Δ1.eviously . EΔ1.1σ7All of these procedures were carried out exactly as previously described .70) or TraR on the energy diagram for transcription initiation using the flux calculator developed by 701.1 deletion  effect on KB  and a large effect on subsequent isomerization rates  was included in the illustrations of the kinetic scheme but did not need to be accounted for in the calculations of fold-inhibition or fold-activation because deletion of σ701.1 or addition of TraR has no effect on this step. In this way, the rate constants for the first step of the flux calculator  as opposed to promoter DNA. The binding of TraR is shown to lead to significant and novel conformational changes in RNAP, which were not evident in prior crystallographic studies, that now account for TraR's function. The structural data, combined with biochemical analyses, provide a mechanism for how TraR binding alters the RNAP conformation and conformational dynamics to modulate the energy landscape of RNAP/promoter formation and transcription initiation. The findings represent a significant step toward enhancing our understanding of the molecular mechanism of how TraR and its homologs, such as DksA, regulate transcription.The current work presents a structural analysis of the transcription factor TraR bound to Decision letter after peer review:E. coli TraR allosterically regulates transcription initiation by altering RNA polymerase conformation and dynamics" for consideration by eLife. Your article has been reviewed by two peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Gisela Storz as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Dong Wang (Reviewer #1).Thank you for submitting your article "The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.Summary:E. coli RNAP and σ-70, with or without TraR, and of RNAP•σ-70 bound to a TraR-repressed promoter are presented. The authors find that upon binding of TraR, substantial conformational changes occur in RNAP that account for TraR's function. These changes are not seen in previous crystallographic analysis, which may have been limited by crystal packing constraints, and indicate that TraR modulates RNAP transcription initiation by binding and altering the conformation and dynamics of the polymerase. TraR-induced transitions are further shown by biochemical analyses to be critical for transcription activation or inhibition, depending on the kinetic features of regulated promoters. Taken together, a detailed mechanism is presented for how binding of TraR alters RNAP conformation and dynamics and to modulate the energy landscape of polymerase-promoter formation and transcription initiation.In the present manuscript, Darst and colleagues use cryo-electron microscopy to determine how TraR, a bacterial transcription factor related to the ppGpp-response factor DksA, regulates RNA polymerase function. Structures of The structural work is of significant interest, as the physical mechanisms through which TraR factor (and other transcription factors which bind directly to RNAP) regulate the kinetics of transcription initiation are poorly understood. The manuscript is clearly written and represents a nice example of the growing power of cryo-EM for the analysis of transcription complexes and their regulation. The work should be of general interest to scientists who work in transcription field; however, several issues need to be addressed before a decision on publication can be made.Essential revisions:1) One issue is that no structure of TraR bound to an initiation complex is provided. The authors mention in the Materials and methods that they were unable to obtain a DNA-bound complex of TraR-RNAP in the cryo-EM. This is unfortunate, as the conformational changes that occur in the presence of DNA will be important for fully defining the structural mechanism of TraR. The lack of a structure is puzzling, since it is shown that TraR is active under the cryo-EM conditions . This raises some concerns that the cryo-EM conditions allow TraR-RNAP interactions and subsequent conformational changes, but that these changes are not congruent with those that are required to support DNA binding. In addition, there is a degree of speculation on this issue in the Discussion that does not seem fully substantied in lieu of a DNA-bound TraR-RNAP complex. Some commentary on this point is needed. It is also recommended that the authors tone down some of the language on this point.2) Based on its blocking of the NTP-entry channel, a TraR-bound complex would be, by definition, a repressed complex for initiation. It is less clear how this fact impacts the interpretation of the kinked bridge helix as a mechanism of repression. A simpler model (related to the one laid out in the last paragraph of the Discussion) may be that activation/repression is dictated solely by the lifetime of RPo in the absence of NTPs. In a potentially interesting synthesis, could it be that the BH clash results in a very low affinity between TraR and RPo, which might also explain the inability to obtain TraR RPo structures? If so, it may be that forming TraR-RPc complexes with DNA at low temperature (4C) prior to freezing for cryo-EM would reveal closed complexes with TraR bound. Here, formation of RPo would have a large negative linkage with TraR binding and formation of RPo would eject TraR in much the way as it ejects σ1.1. This hypothesis could also directly tested by comparing the affinity of TraR to RNAP alone and preformed RNAP RPo with mismatched DNA.3) In multiple instances, the manuscript suggests links between structural observations and kinetics or dynamics. However, these links are hypothetical, as no time-dependent data are presented; the observed conformational heterogeneity (which is well-characterized here) may not necessarily be truly dynamic. This mis-conception is strengthened by wording throughout the paper, exemplified by the following in the Introduction, "…new software tools allow for the analysis of molecular motions in the cryo-EM data…" and from the Abstract, "that quantitative analysis of RNAP conformational heterogeneity revealed TraR-induced changes in RNAP dynamics." While it is accepted that motions are implied by the heterogeneity, the language connecting these two phenomena should be more nuanced.4) The authors state that, "dampening of motion extent is proposed to facilitate the "nucleation of strand opening". But the referenced manuscript  proposes that motions between cRNAP and oRNAP are important for strand opening, stating: "Thus, −11 base-pair perturbations obviate the requirement for cRNAP, confirming that transient clamp closure nucleates melting," and "−10 element recognition by cRNAP nucleates melting, followed by transition into oRNAP form, allowing DNA loading and unwinding in the cleft; the final closure seals the promoter complex." Based on the model presented in Figure 4 of Feklistov, 2017, it would seem that restricting the ability to form oRNAP may activate or inhibit strand separation depending on the kinetics of RP1-RP2-RPo. Please comment.5) There is not a clear synthesis of the variety of inferred structural effects on initiation. How are the effects on the clamp, the bridge helix, and the SLIs integrated to produce activation or inhibition? A kinetic model/schematic that concretely summarizes the kinetic and structural hypotheses for the reader in the context of transitions between different intermediate structures would go a long way to clarifying and synthesizing the proposed TraR model.E. coli TraR allosterically regulates transcription initiation by altering RNA polymerase conformation" for consideration by eLife. Your article has been reviewed by two peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Gisela Storz as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Dong Wang (Reviewer #1).Thank you for submitting your article "The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.ßSummary:There are still a few questions surrounding the work, mostly with respect to the description of a mechanism for activation. These aspects of the manuscript require some attention.Essential revisions:1) It would seem from the response that the authors are preparing a subsequent manuscript that will directly address the issue of the mechanism bound to initiation complexes, and that this may be considerably more complicated than the effects of TraR on the polymerase alone. Given that the paper pertains to the mechanism of TraR-based regulation, it is important to delineate how the present work, in particular the molecular mechanisms in the Discussion, stands on its own in absence of the DNA-bound models. In particular, to what extent will the observations described here be relevant after the second paper comes out?2) The response states that the mechanism of inhibition is due to the trapping of an early bound intermediate on repressed promoters. It is also mentioned that on these promoters, bound complexes are short lived and that TraR will likely be present for the entirety of the time. This seems sensible. However, it is also claimed that "Due to differences in the rate-determining steps at inhibited vs. activated promoters, again described in the Discussion, TraR facilitates rather than inhibits RPo formation at activated promoters. The longer lifetime of the open complex at these promoters results in dissociation of TraR before RNAP dissociates from the promoter, allowing NTP entry and catalysis to proceed." In addition, the response states that "In brief, on activated promoters with long RPo lifetime, TraR would be expected to dissociate from the promoter complex before RNAP would dissociate from the promoter, leaving ample time for activation to occur."The proposed logic for activation is unclear. How can TraR activate transcription via early dissociation? What is "ample time for activation to occur?" After it leaves, one might expect that transcription would be de-repressed to basal levels – not activated. Also, relative to the landscapes shown in Figure 8: these are showing landscapes in the presence and absence of a factor, and factor dissociation during the process is not modeled. It is unclear how the landscapes relate to the hypothesis for activation. Please clarify these points in the manuscript.3) The description of the mechanism via the use of energy landscapes has the potential to greatly clarify the hypotheses made and is a strong addition to the paper. The kinetic/thermodynamic features of the activated and repressed promoters are important should be described in the paper. It seems that one critical component may be the rate of escape. In addition, please provide details on how the flux calculator from Galburt et al. was adapted to a four-state scheme. The on-line calculator appears to only have three states.4) The promoters shown as examples for activated-and-repressed , or as activated-but-no-effect  appear to differ in the rate of promoter escape, which is not mentioned in the manuscript. Could the effect of promoter escape rate revealed by the landscapes be the determinant for TraR-based activation/repression? The landscapes need to be described quantitatively so as to allow others to recapitulate the results. Please comment/clarify.5) There is prior work on the role of different polymerase domains on initiation intermediates  that should probably be considered. Essential revisions:1) One issue is that no structure of TraR bound to an initiation complex is provided. The authors mention in the Materials and methods that they were unable to obtain a DNA-bound complex of TraR-RNAP in the cryo-EM. This is unfortunate, as the conformational changes that occur in the presence of DNA will be important for fully defining the structural mechanism of TraR. The lack of a structure is puzzling, since it is shown that TraR is active under the cryo-EM conditions . This raises some concerns that the cryo-EM conditions allow TraR-RNAP interactions and subsequent conformational changes, but that these changes are not congruent with those that are required to support DNA binding. In addition, there is a degree of speculation on this issue in the Discussion that does not seem fully substantied in lieu of a DNA-bound TraR-RNAP complex. Some commentary on this point is needed. It is also recommended that the authors tone down some of the language on this point.rrnB P1 promoter fragment, as mentioned in the Materials and methods. We have subsequently obtained cryo-EM structures of TraR-RNAP-promoter DNA complexes using the rpsT P2 promoter fragment . Both rrnB P1 and rpsT P2 are inhibited by TraR. The rrnB P1 promoter is known to form a very unstable complex with RNAP with a very short half-life, which is further inhibited by TraR. The rpsT P2 promoter also forms relatively unstable complexes but more stable than rrnB P1 , consistent with our results. Our study of the TraR-RNAP-rpsT P2 complexes has revealed 7 new structures that represent intermediates along the pathway of RPo formation – adequately addressing this work here is way beyond the scope of this manuscript and is the subject of a separate manuscript submitted elsewhere. Nevertheless, the cryo-EM conditions and subsequent conformational changes support DNA binding. Here we focus on conformational changes that TraR induces by binding to RNAP, which provide structure-based hypotheses for the differential effects of TraR on transcription initiation . These hypotheses are then tested with biochemical experiments.We were unable to form a TraR-RNAP-promoter DNA complex with an In response to this and other reviewer comments, we have completely re-written and re-organized the Discussion section.2) Based on its blocking of the NTP-entry channel, a TraR-bound complex would be, by definition, a repressed complex for initiation. It is less clear how this fact impacts the interpretation of the kinked bridge helix as a mechanism of repression. A simpler model (related to the one laid out in the last paragraph of the Discussion) may be that activation/repression is dictated solely by the lifetime of RPo in the absence of NTPs. In a potentially interesting synthesis, could it be that the BH clash results in a very low affinity between TraR and RPo, which might also explain the inability to obtain TraR RPo structures? If so, it may be that forming TraR-RPc complexes with DNA at low temperature (4C) prior to freezing for cryo-EM would reveal closed complexes with TraR bound. Here, formation of RPo would have a large negative linkage with TraR binding and formation of RPo would eject TraR in much the way as it ejects σ1.1. This hypothesis could also directly tested by comparing the affinity of TraR to RNAP alone and preformed RNAP RPo with mismatched DNA.The reviewer raises several interesting and important questions that are relevant to the complexity of the regulatory mechanism employed by TraR. We revised the Discussion, especially the sections entitled “Structural Mechanisms of Activation by TraR” and “Structural Mechanisms of Inhibition by TraR”, to address these questions and to describe our model more comprehensively. We also address these issues by pointing out specific data presented here, as well as in previous publications, that support our model, and we added a new figure  to summarize features of the model.Specifically, we address the questions raised as follows:‘Based on its blocking of the NTP-entry channel, a TraR-bound complex would be, by definition, a repressed complex for initiation.'2a) Yes, we agree that a TraR-bound complex would be repressed for initiation, and this would be true whether the promoter is a negatively or positively regulated one. Thus, this cannot be the mechanism for activation. For the following reasons  we do not think that blocking of NTP entry is the primary mechanism for inhibition.Due to the TraR- induced conformational changes in RNAP (detailed in our paper) that severely reduce RPo formation by stabilizing an earlier intermediate on the pathway and by weakening duplex DNA interactions at inhibited promoters, transcription initiation is blocked before the stage of NTP entry and catalysis. Footprints of negatively-regulated promoter complexes containing TraR display protection from DNaseI characteristic of earlier intermediates rather than RPo, consistent with this interpretation.rrnB P1, which form extraordinarily short-lived promoter complexes with RNAP, the lifetime of the factor with RNAP is longer than the lifetime of RNAP with the promoter.Our earlier data  showed that DksA  has 10-fold reduced binding affinity for RPo relative to RNAP holoenzyme. Consistent with these data, recent single molecule fluorescence data from Gelles and colleagues  showed that secondary channel binding factors  bind to RNAP to form an RNAP-factor complex, before RNAP binds to the promoter, and the factors remain bound to RNAP on the promoter until RNAP dissociates. On negatively-regulated promoters like Due to differences in the rate-determining steps at inhibited vs. activated promoters, again described in the Discussion, TraR facilitates rather than inhibits RPo formation at activated promoters. The longer lifetime of the open complex at these promoters results in dissociation of TraR before RNAP dissociates from the promoter, allowing NTP entry and catalysis to proceed.'It is less clear how this fact impacts the interpretation of the kinked bridge helix as a mechanism of repression. A simpler model (related to the one laid out in the last paragraph of the Discussion) may be that activation/repression is dictated solely by the lifetime of RPo in the absence of NTPs.'2b) We agree with the reviewer that the lifetime of RPo is an important determinant of whether a promoter is activated or repressed, as described above, but we think that it is part of a more complex multistep mechanism. We propose that kinking of the bridge helix (BH) plays a role in the mechanism of inhibition by reducing the stability and occupancy of RPo by preventing a template strand interaction with the BH. In addition, the template strand-BH interaction correctly positions the template strand in the active site, and a BH kink would therefore inhibit catalysis. We suggest while most RNAP-TraR complexes at an inhibited promoter would not proceed to forming an open complex, the small fraction that do would be inhibited by the BH kink and restriction of NTP access, thus ensuring efficient inhibition.‘In a potentially interesting synthesis, could it be that the BH clash results in a very low affinity between TraR and RPo, which might also explain the inability to obtain TraR RPo structures?'2c) Yes, this idea is supported by our previous observation of a 10-fold reduced affinity of DksA, a TraR homolog, for a pre-formed RPo . Since TraR has a similar affinity for RNAP , we propose that TraR also has a reduced affinity for pre-formed RPo. We suggest that RPo formation can occur in a small fraction of complexes in the presence of bound TraR, and that blocking catalysis would be part of a multipartite mechanism that would ensure efficient inhibition of this small fraction of complexes.'If so, it may be that forming TraR-RPc complexes with DNA at low temperature (4C) prior to freezing for cryo-EM would reveal closed complexes with TraR bound. Here, formation of RPo would have a large negative linkage with TraR binding and formation of RPo would eject TraR in much the way as it ejects σ1.1. This hypothesis could also directly tested by comparing the affinity of TraR to RNAP alone and preformed RNAP RPo with mismatched DNA.'2d) Several different TraR-RNAP-promoter complexes formed with DNA templates containing or lacking mismatches have been analyzed using single particle cryo EM. These results identify the structures of several different intermediates in the RPo formation pathway. Their description is beyond the scope of the present manuscript and is being reported elsewhere.3) In multiple instances, the manuscript suggests links between structural observations and kinetics or dynamics. However, these links are hypothetical, as no time-dependent data are presented; the observed conformational heterogeneity (which is well-characterized here) may not necessarily be truly dynamic. This mis-conception is strengthened by wording throughout the paper, exemplified by the following in the Introduction, "…new software tools allow for the analysis of molecular motions in the cryo-EM data…" and from the Abstract, "that quantitative analysis of RNAP conformational heterogeneity revealed TraR-induced changes in RNAP dynamics." While it is accepted that motions are implied by the heterogeneity, the language connecting these two phenomena should be more nuanced.There are two types of conformational heterogeneity characterized by cryo-EM in this study – we can call them digital and analog:70(I) and TraR-Eσ70(II), where the two consensus structures differ in the disposition of β'Si3, which is seen in two discrete positions .I) Digital – The complexes exist in relatively discrete conformational states that can be parsed using the software tools in RELION. These are represented here by TraR-Eσ70(I) and TraR-Eσ70(II)], the RNAP clamp is also conformationally heterogeneous but does not parse into discrete bins. Rather, the clamp positions are characterized by continuous ~Gaussian distributions .II) Analog – Within these classes of particles Essential revisions:1) It would seem from the response that the authors are preparing a subsequent manuscript that will directly address the issue of the mechanism bound to initiation complexes, and that this may be considerably more complicated than the effects of TraR on the polymerase alone. Given that the paper pertains to the mechanism of TraR-based regulation, it is important to delineate how the present work, in particular the molecular mechanisms in the Discussion, stands on its own in absence of the DNA-bound models. In particular, to what extent will the observations described here be relevant after the second paper comes out?eLife submission.The current paper focuses on the structural mechanisms for both inhibition and activation by TraR. The second paper uses TraR as a tool to populate promoter melting intermediates with an inhibited promoter. The subject of this second paper is not TraR, nor its mechanism. The second paper describes the structures of these intermediates and also presents structure-based biochemical tests to support our proposal that these intermediates describe the normal promoter melting pathway even in the absence of TraR. Thus, the subject of the second paper is promoter melting, not TraR mechanism. There are no results in the second paper that contradict or supersede the results of this 2) The response states that the mechanism of inhibition is due to the trapping of an early bound intermediate on repressed promoters. It is also mentioned that on these promoters, bound complexes are short lived and that TraR will likely be present for the entirety of the time. This seems sensible. However, it is also claimed that "Due to differences in the rate-determining steps at inhibited vs. activated promoters, again described in the Discussion, TraR facilitates rather than inhibits RPo formation at activated promoters. The longer lifetime of the open complex at these promoters results in dissociation of TraR before RNAP dissociates from the promoter, allowing NTP entry and catalysis to proceed." In addition, the response states that "In brief, on activated promoters with long RPo lifetime, TraR would be expected to dissociate from the promoter complex before RNAP would dissociate from the promoter, leaving ample time for activation to occur."The proposed logic for activation is unclear. How can TraR activate transcription via early dissociation? What is "ample time for activation to occur?" After it leaves, one might expect that transcription would be de-repressed to basal levels – not activated. Also, relative to the landscapes shown in Figure 8: these are showing landscapes in the presence and absence of a factor, and factor dissociation during the process is not modeled. It is unclear how the landscapes relate to the hypothesis for activation. Please clarify these points in the manuscript.We have further revised the Discussion section on “Structural mechanism for TraR-mediated activation” to clarify the model and address the questions raised by the reviewer. The first question concerned how dissociation of TraR from the open complex (RPo) could activate transcription. Confusion about this subject could have arisen because we previously addressed it at the end of the Discussion section on “Structural Mechanism of TraR-mediated inhibition” instead of in the section on “Structural mechanism of activation”. We have now moved the explanation to the activation section so that all features of the proposed model for activation are now presented together in one section.1.1 in the main DNA binding channel , thereby increasing the ability of promoter DNA to displace σ1.1 from this channel. TraR also alters the position of β’Si3, contributing to activation . Contrary to what is implied in the comment, these enhancements to the overall rate occur while TraR is still present in the complex – the increase in the rate and amount of RPo formed while TraR was present leads to the increase in transcription once TraR dissociates.In summary, we have previously published data showing that TraR and its homolog DksA enhance the transcription output from activated promoters relative to that with RNAP alone , and that RPo at activated promoters is very stable to competitor challenge . In addition, kinetic analyses showed that the presence of DksA enhanced the rate of formation and the amount of competitor stable RPo formed at an activated promoter relative to that formed with RNAP alone . In the present paper we present structural and biochemical evidence that these effects occur primarily because TraR-induces conformational changes in the β subunit that allosterically alter the position of σSince the structural data shown here indicate that TraR bound to the complex would sterically block iNTP access to the active site, t-strand positioning and catalysis , TraR must therefore dissociate from the complex in order for transcription to initiate from the activated promoter. The proposal that TraR dissociates from RPo at an activated promoter prior to dissociation of RNAP from the promoter is consistent with previously published data. The lifetimes of RPo at activated promoters  are much longer than that of the transcription factors on RNAP .A second part of the reviewer’s comments concerns whether TraR “de-represses basal activity” rather than really activates transcription. We refer to “basal” activity as the activity observed with RNAP alone in the absence of TraR. We have shown previously that TraR increases transcription 4-6 fold over the amount observed without TraR, i.e. it really activates . We have revised a paragraph  to clarify what is meant by basal transcription.We have also included a description in the revised Discussion section about how the energy landscape shows the activation step. We have also modified the kinetic scheme at the top of Figure 8 to reflect TraR (T) dissociation that is required for initiation of RNA chain synthesis and transition of RPo to RPitc.3) The description of the mechanism via the use of energy landscapes has the potential to greatly clarify the hypotheses made and is a strong addition to the paper. The kinetic/thermodynamic features of the activated and repressed promoters are important should be described in the paper. It seems that one critical component may be the rate of escape. In addition, please provide details on how the flux calculator from Galburt et al. was adapted to a four-state scheme. The on-line calculator appears to only have three states.The general kinetic/thermodynamic features of inhibited vs. activated promoters are discussed in the Discussion. When the energy diagrams are first introduced , we have added a note that the features of inhibited vs. activated promoters are described later in the Discussion. We have also added new references establishing the 3-step minimal linear mechanism for RPo formation (or 4-step mechanism if one includes promoter escape). Basically, complexes at inhibited promoters pass through the pathway rapidly but the final RPo is relatively unstable; in the absence of high NTP concentrations, RPo exists in equilibrium with earlier intermediates. By contrast, activated promoters form RPo slowly, but the RPo that is formed is very stable. In terms of the basic features of the energy diagram, inhibited promoters have low kinetic barriers  but the valleys going from left to right are generally uphill. Activated promoters have high kinetic barriers  but the valleys going from left to right are generally downhill. We have tried to capture these basic features in the basal energy diagrams (the black lines) in Figure 4—figure supplement 1C and Figure 8 (these figures have been revised to better illustrate these points).Regarding the rate of escape, there is no evidence that promoter escape is rate-limiting or that TraR alters the rate of escape from either the activated or inhibited promoters used here. Our models for activation and inhibition propose that TraR would not be associated with the final RPo at the activated promoters (see above response to comment 2), so would not affect the escape rate. For inhibited promoters, TraR depopulates RPo by stabilizing an earlier intermediate complex, thereby preventing initiation. Thus, it is unlikely that TraR remains bound to RPo except in a small fraction of the population in which TraR binding would be expected to sterically occlude catalysis .The details of how we used the flux calculator of Galburt, 2018, are now described in Materials and methods . The input parameters and results of the flux calculator calculations are now tabulated in Supplementary file 2.4) The promoters shown as examples for activated-and-repressed , or as activated-but-no-effect  appear to differ in the rate of promoter escape, which is not mentioned in the manuscript. Could the effect of promoter escape rate revealed by the landscapes be the determinant for TraR-based activation/repression? The landscapes need to be described quantitatively so as to allow others to recapitulate the results. Please comment/clarify.The details of how we used the flux calculator of Galburt, 2018, are now described in Materials and methods . The input parameters and results of the flux calculator calculations are now tabulated in Supplementary file 2. The rates of escape for the inhibited promoter in Figure 4—figure supplement 1C and in Figure 8, and the activated promoter in the two cases, are the same, and these are not altered by TraR in our model.5) There is prior work on the role of different polymerase domains on initiation intermediates  that should probably be considered.1.1 on promoter complex formation and stability. Their results, specifically that deletion of σ1.1 increases RPo formation at some promoters and decreases it at others, are consistent with effects of the σ1.1 deletion on transcription observed here at activated or inhibited promoters . Work from the Record lab also showed a role for Si3 in stabilization of RPo . This is consistent with our finding that Si3 plays a role in promoter activation by TraR  and suggests that Si3 could affect activation by altering complex stability. This is now mentioned in our revised Discussion section (see the fourth paragraph of the aforementioned subsection).We agree with the reviewer that previous work about the role of different RNAP domains should be mentioned. We have included a sentence  describing conclusions from previous work from the Record and Hinton labs on effects of the deletion of σ


Helicobacter pylori the expression of chaperone-encoding operons is under control of two auto-regulated transcriptional repressors, HrcA and HspR, with the latter acting as the master regulator of the regulatory circuit. To further characterize the HspR regulon in H. pylori, we used global transcriptome analysis (RNA-sequencing) in combination with Chromatin Immunoprecipitation coupled with deep sequencing (ChIP-sequencing) of HspR genomic binding sites. Intriguingly, these analyses showed that HspR is involved in the regulation of different crucial cellular functions through a limited number of genomic binding sites. Moreover, we further characterized HspR-DNA interactions through hydroxyl-radical footprinting assays. This analysis in combination with a nucleotide sequence alignment of HspR binding sites, revealed a peculiar pattern of DNA protection and highlighted sequence conservation with the HAIR motif (an HspR-associated inverted repeat of Streptomyces spp.). Site-directed mutagenesis demonstrated that the HAIR motif is fundamental for HspR binding and that additional nucleotide determinants flanking the HAIR motif are required for complete binding of HspR to its operator sequence spanning over 70 bp of DNA. This finding is compatible with a model in which possibly a dimer of HspR recognizes the HAIR motif overlapping its promoter for binding and in turn cooperatively recruits two additional dimers on both sides of the HAIR motif.The ability of pathogens to perceive environmental conditions and modulate gene expression accordingly is a crucial feature for bacterial survival. In this respect, the heat-shock response, a universal cellular response, allows cells to adapt to hostile environmental conditions and to survive during stress. In the major human pathogen  Helicobacter pylori represents one of the most widespread human pathogens, recognized as the principal causative agent of different gastrointestinal severe diseases such as atrophic gastritis, peptic ulcer, MALT-lymphoma and gastric adenocarcinoma , characterized by a relative low abundance of genes encoding regulators of transcription. In this respect, it has been speculated that this paucity of transcriptional regulators could reflect the adaptation of H. pylori to its very restricted niche in the mucus layer of the human stomach and could be linked to the lack of competition from other microorganisms . The main function of HSPs is to assist the folding of newly synthetized polypeptides, as well as assembly, transport and degradation of cellular proteins under both normal and adverse growth condition   . Also inene clpB . In H. pIR motif . While iepressor . The ide. pylori  and in s. pylori . Moreoveaperones .H. pylori HspR to the heat-shock regulon. While the heat-shock regulon includes many genes with key cellular functions, HspR only binds to a limited number of genomic sites. Furthermore, by means of the high-resolution hydroxyl-radical footprinting technique, we further characterized the HspR-DNA interactions at the molecular level. The data provide a more detailed comprehension of the interaction between HspR and its target DNA sequences and, at least on its own promoter, let us to propose a cooperative DNA-binding mechanism of three HspR dimers per operator sequence.In the present study, by combining ChIP-sequencing and RNA-sequencing we investigated more in detail the direct or indirect contribution of Helicobacter pylori strains (Table 1) were recovered from frozen glycerol stocks on Brucella broth agar plates, containing 5% fetal calf serum (FCS), in a 9% CO2-91% air atmosphere at 37°C and 95% humidity in a water jacketed incubator (Thermo Scientific). Liquid cultures were grown in Brucella Broth supplemented with 5% FCS with gentle agitation (120 rpm). E. coli strains DH5α and BL21 (DE3) (Table 1) were grown on Luria-Bertani (LB) agar plates or LB liquid broth with vigorous agitation (250 rpm); when required, ampicillin was added to the medium to achieve a final concentration of 100 μg/ml.Helicobacter pylori strains (Table 1) were grown with gentle agitation (120 rpm) in 30 ml of Brucella broth at 37°C until mid-exponential phase (OD = 0.7). For heat-shock treatment, the wild type (WT) culture was split into 15 ml-aliquots and one sample was subjected to heat-shock at 42°C for 30 min . A volume of 10 ml cell culture was then added to 1.25 ml of ice-cold EtOH-phenol stop solution  to stop growth and prevent RNA degradation. Cells were pelleted, stored at −80°C, and then used to extract total RNA with TRI-reagent (Sigma-Aldrich), according to manufacturer’s protocol.TM v2 RNAseq library preparation kit  starting from 50 ng of previously rRNA-depleted RNA from each biological replicate and for all the conditions analyzed. Then, each library was multiplexed in equal amounts and sequenced on a GAIIX Illumina sequencer and 85 bp reads were produced. A minimum of 7 Million reads were obtained for each of the samples and for each replica. Bowtie 2 (v2.2.6)  and alignment score >−15; for multi-mapping reads alignment score was set ≥−15. H. pylori G27 RefSeq annotation (GCF_000021165.1) in the version released on sept-2017 was used as the reference for gene annotation to which we manually added validated ncRNAs (Supplementary Table S1). We also revised the annotation of protein coding genes that, based on our sequencing data, were improperly annotated as pseudogenes in this version of the reference genome , indicating them as “protein-coding∗” in Supplementary Table S1. BEDTools (v2.20.1) (Supplementary Table S2). This analysis revealed that 99% of the transcripts were covered by at least one strand specific read and a minimum of 46 reads were counted on 90% of them. The R package DESeq2 (v1.4.5)  lower than 0.01 and log2 fold changes (log2FC) > |1| . Raw data are publicly available at Sequence Reads Archive under accession number BioProject PRJNA421261.Ribosomal RNAs were depleted starting from 1 μg of total RNA from each of the conditions analyzed by using the RiboZero Gram negative kit . Strand specific RNA-seq libraries were prepared by using the ScriptSeq(v2.2.6)  was usedd ncRNAs  (highligv2.20.1)  and SAMtv2.20.1)  were use(v1.4.5)  was thenSupplementary Table S1).To evaluate functional enrichments in the DEGs lists, we retrieved COG functional classes for all the protein coding genes present in our annotation file through the NCBI CDD database . We obta2SO4 and subsequent resuspension in water. H. pylori G27 wild type and hspR mutant strains were liquid-grown to an OD600 of 0.7, crosslinked, sonicated and immunoprecipitated as previously described  and IP (IP wt) samples sequencing on the H. pylori G27 genome. End-to-end mapping was performed and non-deterministic option was specified. High quality reads were then selected requiring: for uniquely mapping reads MAPQ > 30  and alignment score > −10 while, for multi-mapping reads, the alignment score was set ≥−10. On average, more than 98% of them mapped on the H. pylori G27 reference genome. The ChIP-seq data quality was evaluated by using ENCODE quality metrics1 and the values obtained are provided in Supplementary Table S2. To perform peak calling, the Homer (v4.7.2)  read density 4-fold greater than in the surrounding 10 kb region; (3) the ratio between the number of unique positions containing reads in the peak and the expected number of unique positions given the total number of reads in the peak lower than 2. Only the peaks identified in both biological replicates and having overlapping genomic coordinates were considered significantly reliable and included in the final peak list. The peaks having their center within −100/ + 30 bp from a transcription start site (TSS) were defined as promotorial, while the remaining peaks were divided in intragenic, when their center was mapping within a predicted coding region, or intergenic, when it was mapping outside from annotated regions . TSS were identified first by blasting 50 bp upstream of each of the transcription initiation sites reported by PRJNA421261.Illumina libraries were prepared following the Illumina TruSeq ChIP-seq DNA sample preparation protocol starting from 5 ng of immunoprecipitated-DNA for each of the strains and each of the two biological replicates. Each library was sequenced on a GAIIx or MiSeq Illumina sequencer and 51 bp single stranded reads were produced. At least 2 Million of raw reads were obtained for each IP sample and biological replicate. Bowtie 2 (v2.2.6)  was used(v4.7.2)  algorithDNA manipulations were performed as described by 6-tagged recombinant HspR protein was overexpressed in E. coli BL21 (DE3) cells and affinity purified as previously described  avoiding any trace of glycerol, prior to the DNA binding experiment, and stored at −80°C. Protein concentration was determined by Bradford colorimetric assay (BioRad) and purity assayed by SDS-PAGE.Hisescribed . The purH. pylori G27 encompassing HspR binding sites on hrcA, groES, cbpA promoters and speA coding region were PCR amplified with specific primers (Table 2) and cloned into the pGEM-T-Easy plasmid (Table 1). The M1, M3, M4 and M5 Pcbp mutant probes were generated by annealing complementary oligonucleotides to form a double stranded DNA fragment with compatible overhangs required to clone it in the pGEM-T-Easy plasmid previously digested with the appropriate restriction enzymes. The M2 and M1+2 Pcbp mutant probes were generated through site-directed mutagenesis using the plasmid pGEM-T-Easy, harboring the Pcbp wild type sequence, as DNA template and primers listed in Table 2.Genomic regions of 32 P]-ATP and T4 polynucleotide kinase and gel purified. Hydroxyl-radical footprinting experiments were performed as previously described  for 15 min at room temperature, including 200 ng of sonicated salmon sperm DNA as a non-specific competitor in a final volume of 30 μl. Partial digestions of the labeled probes were achieved using 2 μl each of the following solutions: 125 mM Fe (NH4)2(SO4)2 250 mM EDTA, 1% H2O2 and 100 mM DTT. After 2 min incubation, the reaction was quenched by the addition of 25 μl of OH Stop Buffer . Samples were phenol/chloroform extracted, ethanol precipitated and resuspended in 12 μl of Formamide Loading Buffer. Next, samples were denatured at 100°C for 5 min, separated on a 8M urea-8.4% polyacrylamide sequencing gel in TBE buffer and autoradiographed.Probe DNA fragments obtained by digestion with the appropriate restriction enzymes were 5′ end-labeled with [γescribed  with somcbp and mutated Pcbp M1 + 2 promoter regions were PCR amplified with specific primers listed in Table 2, using as template the H. pylori G27 genomic DNA and the plasmid pGEM-T-Easy-HBScbp M1 + 2 (Table 1), respectively. Then, the generated DNA fragments were digested with appropriate restriction enzymes and cloned into the pVAC plasmid (Table 1). The luxC gene was PCR amplified from the pVCC vector (Table 1) and cloned downstream the Pcbp promoters into the pVAC vector. These plasmids were used to transform the H. pylori G27 wild type acceptor strain in the vacA locus. The chloramphenicol-selected mutant strains were expanded and the correct insertion was confirmed by PCR using oligonucleotides pair cys-F/vacA-R as primers (Table 2). The H. pylori Pcbp-lux, hspR::Km reporter strains (Table 1) were generated transforming the H. pylori vacA::Pcbp wt-lux and the H. pylori vacA::PcbpM1 + 2-lux acceptor strains (Table 1) with the plasmid pGEM3-hspR::Km (Table 1). The kanamycin-selected mutant strains were expanded and the correct insertion was confirmed by PCR using oligonucleotides pair cbpRTRev/hspRC as primers (Table 2).The wild type PTable 2) at 400 nM concentration in a final volume of 10 μl. qRT-PCR experiments were performed using the following cycling protocol: 95°C for 2 min, then 40 cycles consisting of a denaturation step for 5 s at 95°C followed by 30 s at 60°C . Data were analyzed using the ΔΔCt method, using the housekeeping ppk gene, known to be constitutively expressed, as internal reference, using oligonucleotides ppk RT-F and ppk RT-R as primers (Table 2)  .H. pylori G27 strain and of ΔhspR mutant both grown to the exponential growth phase at 37°C and of the wild type strain subjected to 30 min heat-shock at 42°C .To define the HspR contribution to the heat-shock response, we performed a strand-specific whole transcriptome analysis of the wild type ΔhspR mutant to that of the H. pylori wild type strain, both grown at 37°C (ΔhspR_vs_WT). This analysis showed a total of 65 deregulated genes (log2FC > |1| padj < 0.01) upon hspR gene deletion. Of these, 21 were down-regulated and 44 were up-regulated genes . As expected, among the up-regulated genes we found groES, groEL, grpE, dnaK, and cbpA (padj = 0.01) ΔhspR de-repressed genes , and hrcA transcriptional regulator . As expected, because of the hspR genomic deletion, this gene appeared as down-regulated. The rarA gene, mapping downstream of the hspR gene, possibly due a polar effect of the deletion mutant also appeared as a down-regulated gene. Several transposase coding genes, producing the “Mobilome: prophages, transposons” category enrichment (padj = 0.00009), and several genes involved in the “Inorganic ion transport and metabolism” were also down-regulated.We defined the role of HspR in standard growth conditions by comparing the transcriptome of the and cbpA , which sH. pylori wild type G27 strain showed a total of 134 differentially expressed genes (log2FC > 1 padj < 0.01) when compared to the wild type sample not subjected to heat-shock (HS_vs_WT). Of these genes, 83 were up-regulated and 51 were down-regulated . Functional annotation and enrichment analysis  revealed that up-regulated genes were enriched in “Post-translational modification, protein turnover, chaperones” (padj = 0.0007) and “Mobilome: prophages, transposons” class . Several of the 51 down-regulated genes were also annotated to “Cell wall/membrane/envelope biogenesis” and “Translation, ribosomal structure and biogenesis” categories  without statistical significance.Upon heat-shock at 42°C the transcriptome of the hspR deletion mutant (65 deregulated genes) we identified 25 genes that were deregulated in both datasets . Among these genes, 10 were oppositely regulated and 14 were similarly regulated: 2 down-regulated and 12 up-regulated . The similarly up-regulated genes included all the genes of the operons already known to be directly repressed by HspR and induced by heat-shock . The remaining eleven genes were induced by heat-shock and repressed in the hspR mutant strain, thus showing opposite behavior. Considering that HspR acts as a repressor factor, these genes are probably indirectly controlled by HspR.Comparing the transcriptome of the heat-shock response (134 deregulated genes) to that of the at-shock . In addiOverall, this analysis highlights that HspR could be involved in the control of maximum 18% (25 out of 134) of the genes whose transcription is heat responsive and that it represses the transcription of only 14 genes.in vivo by HspR, therefore, the genes directly controlled by HspR binding, we performed a Chromatin Immunoprecipitation assays followed by deep sequencing (ChIP-seq) in H. pylori G27 wild type and ΔhspR strains . To identify HspR bound regions (peaks), ChIP-seq signals obtained from each wild type sample (IP wt) were compared to those resulting from the pool of ΔhspR mutant samples (IP ΔhspR) . Only the significant peaks in both replicates were considered in the final peak list. Surprisingly, this analysis identified only four reproducible HspR binding regions , which were annotated with respect to the latest genome annotation of the H. pylori G27 strain (GCF_000021165.1). According to this annotation, three peaks were classified as promotorial and confirmed HspR direct control for 8 genes of the groES-groEL, hrcA-grpE-dnaK and cbpA-hspR-rarA operons. The remaining peak, mapping inside the coding region of speA gene, was classified as intragenic. Thus, these data suggest that the HspR direct regulon is very restricted and limited to the three multicistronic heat-shock operons. Moreover, through ChIP-seq analysis we confirmed in vivo also the existence of at least one intracistronic binding site, apparently not associated to transcriptional regulation.To identify the genomic regions bound H. pylori HspR binds directly to extended DNA regions of the promoters of the three heat-shock operons  and on the 3′ region of the speA coding sequence. According to previous observations  on the Pcbp, Pgro, Phrc promoters and on the speA coding region. Notably, in all the operators we identify one inverted repeat  similar to the HAIR motif proposed as a consensus sequence for the HspR protein of S.coelicolor (CTTGAGT-N7-ACTCAAG) (Streptomyces spp.Previous DNase I footprinting assays showed that  operons . In partrvations  the protACTCAAG) . It is wcbp region is reported in Figure 4A. As shown in Figure 4B, mutation of one or both arms  of the HAIR-like sequence completely abolished HspR binding to the DNA in the concentration range tested. In particular, partial (M1 and M2) or total substitution (M1 + 2) of the HAIR-like sequence prevented HspR binding to both, the mutagenized sequence  and the flanking upstream and downstream regions of the HAIR-like motif. It is worth mentioning that regions protected in hydroxyl-radical footprinting experiments reflect limited accessibility of radical ions to the DNA minor groove and, for this reason, these protected regions do not necessarily represent the portions of the probe directly contacted by the HspR protein, but regions surrounding short stretches of contacted nucleotides. Considering that HspR footprint regions surround the conserved HAIR-like element , our data suggest that HspR could interact with the HAIR-like element in the DNA major groove narrowing the adjacent minor grooves, which results in the protection observed in vitro by hydroxyl-radical footprinting. Furthermore, mutation of the non-conserved sequence in between the two inverted repeats  showed a barely detectable protection of HspR only upon addition of high amount of the protein to the reaction . This is likely due to a significant loss of protein affinity to the DNA, suggesting that, besides the HAIR-like motif, also this non-conserved DNA element is important for HspR binding. These data show for the first time that in H. pylori the HAIR-like sequence is an essential DNA element for the specific binding of HspR to the Pcbp promoter region. Likely, HspR binding to the operator is driven by a specific recognition of determinants within the HAIR-like sequence, including the non-conserved spacer between the inverted repeat.To ascertain the functional importance of the HAIR-like sequence elements in HspR binding to DNA, we decided to introduce bases substitutions in the inverted repeat and monitor their effects on HspR DNA-binding through hydroxyl-radical footprinting assays. A schematic representation of the wild type and mutant sequences of the HspR binding site on the Pin vivo the functional significance of the central HAIR-like motif, we generated a reporter construct in which the wild type or HAIR-like mutant Pcbp promoter (PcbpM1 + 2) was fused upstream of a 5′ fragment of the luxCDABE reporter cassette and, upon integration in H. pylori chromosome, transcript level was assayed (through RT–qPCR with luxC specific primers) during exponential growth . When the central HAIR-like inverted repeat was mutated in the G27 wild type strain, a consistent increase in the amount of transcript from the PcbpM1 + 2 promoter with respect to the wild type Pcbp promoter was observed . The about 6.5-fold increase of transcript from the PcbpM1 + 2 promoter is in line with the previously observed de-repression of the Pcbp promoter in the H. pylorihspR-mutant strain . These in vivo data are consistent with the in vitro observations, indicating that the central HAIR-like motif drives specific binding of HspR to its operator on Pcbp with concomitant promoter transcriptional repression.To characterize t strain . Accordicbp promoter and assayed for HspR binding by hydroxyl-radical footprintings. Mutations were introduced in the non-conserved spacer region between two 4-bp protected tracts on both sides of the inverted repeat . Surprisingly, as shown in Figure 6B, the addition of increasing amounts of HspR to the mutant probes led to the disappearance of regions of protection (marked in light gray) only on the side of the mutated region, while it was unaffected on both the inverted repeat of the HAIR-like sequence and on the opposite side of the mutation . In conclusion, these data support the pivotal role of the HAIR-like motif for HspR-DNA binding in H. pylori and demonstrate that also other non-conserved DNA regions surrounding the HAIR-like motif are important elements that allow HspR to completely occupy its extended binding sites.In order to understand if DNA sequences flanking the HAIR-like motif are important for HspR-DNA recognition and binding, we designed two mutant probes of the PH. pylori their expression level is tightly modulated by the concerted action of two transcriptional repressors, HrcA and HspR, with this latter acting as the master regulator of this circuit . Accordingly, the heat-shock operons groES-groEL, hrcA-grpE-dnaK and cbpA-hspR-rarA, coding for the major chaperones and heat-shock proteins of H. pylori were clearly up-regulated. With the exception of the hspR and rarA genes, these operons, known to be directly repressed by HspR . In contrast, ChIP-seq analysis showed only four already known in vivo HspR genomic binding sites, three of which are associated to the promoter regions of the heat-shock operons and one mapping within the coding sequence of the speA gene , surprisingly, the transcription of genes coding for stress related proteases seems to be unaffected by the heat challenge. Furthermore, while in some other bacteria, like for example Campylobacter jejuni and S. coelicolor, several protease-encoding genes have been shown to belong to the HspR regulon . Binding of HspR to promoter regions shows 7 short protected DNA tracts spaced by non-protected regions of 7/8 nucleotides, while binding within the speA coding sequence appears at higher protein concentrations and shows only 5 short protected regions. This different binding pattern could be related to the fact that the latter binding site on the coding sequence of speA gene appears to be not associated to transcriptional regulation. In fact, RNA-seq analysis revealed no changes in the level of speA transcript in the hspR mutant strain, nor of neighboring genes. This is not surprising, as the advent of the “omics” era highlighted binding of regulatory proteins to a number of binding sites not associated to regulation, such as the H. pylori Fur repressor . These HAIR-like motifs map in the central position of the HspR binding sites on the three heat-shock operons’ promoters and appear to be an essential DNA element for specificity of protein binding. In fact, mutation of one or both arms of this inverted repeat completely abolished the HspR binding to the operator  and prevented in vivo HspR-dependent repression of Pcbp , demonstrating for the first time that the HAIR-like sequence is functionally important also in H. pylori. The DNA-binding mechanism of HspR on its target operators is unique among H. pylori transcriptional regulators characterized so far. Well-studied H. pylori regulatory proteins, such as HP1043, HrcA, NikR and Fur appear to recognize conserved sequence motifs as dimers and protect limited DNA regions  in the promoter region of its DNA targets  and flanking  the HAIR-like motif are important elements for DNA recognition and binding of HspR to the operator. Intriguingly, mutation of one of these elements impaired binding of HspR on the side of the mutation and showed no effects on the binding on the HAIR-like motif on the opposite side of the mutation . This finding is compatible with a cooperative mechanism of HspR binding. Possibly, a dimer of HspR recognizes and binds to the HAIR-like sequence and this in turn drives binding of additional dimers on both sides of the HAIR-like motif . Evidences that HspR could bind its DNA sequences as a dimer have been shown in previous studies, in which it was demonstrated that homologs of H. pylori HspR repressor protein could exists in a dynamic state between the dimeric and monomeric forms in solution , it can also be speculated that HspR DNA binding could be driven by sequence-dependent local shape variations rather than by a base readout mechanism, or by a combination of these two mechanisms. In other words, the crucial role of the HAIR-like motifs for HspR binding could be explained by taking into account local DNA shape variations imposed by a peculiar succession of purine/pyrimidine, rather than considering only the unique chemical signatures of the individual DNA bases. This behavior is common among regulators of MerR family, to which the H. pylori HspR belongs to, employing indirect (shape) readout as part of their DNA binding mechanisms. These considerations could help also to explain our data on the importance of the central region of the HAIR-like motif and of the non-conserved regions on both sides of the HAIR-like motif .Previous DNase I footprinting assays of HspR showed protection of large DNA regions of about 70 bp upstream of the promoters controlling transcription of three heat-shock operons . To deepepressor , the E. e enzyme . Howeverces spp.  (Figure  regions . In the  regions . Howeverg assays . This pe targets , while isolution . MoreoveAD, CP, DR, and VS conceived and designed the experiments. SPE, EP, EF, TB, CP, and AV performed the experiments. SPE, EP, and SPU carried out the data analysis. DR and VS wrote the paper with contributors CP and EP. All the authors reviewed the manuscript.The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.


Gene regulation by enhancers is important for precise temporal and spatial gene expression. Enhancers can drive gene expression regardless of their location, orientation or distance from the promoter. Changes in chromatin conformation and chromatin looping occur to bring the promoter and enhancers into close proximity. αA-crystallin ranks among one of the most abundantly expressed genes and proteins in the mammalian lens. The αA-crystallin locus is characterized by a 16 kb chromatin domain marked by two distal enhancers, 5′ DCR1 and 3′ DCR3. Here we used chromatin conformation capture (3C) analysis and transgenic approaches to analyze temporal control of the mouse αA-crystallin gene. We find that DCR1 is necessary, but not sufficient alone to drive expression at E10.5 in the mouse lens pit. Chromatin looping revealed interaction between the promoter and the region 3′ to DCR1, identifying a novel enhancer region in the αA-crystallin locus. We determined that this novel enhancer region, DCR1S, recapitulates the temporal control by DCR1. Acting as shadow enhancers, DCR1 and DCR1S are able to control expression in the lens vesicle at E11.5. It remains to be elucidated however, which region of the αA-crystallin locus is responsible for expression in the lens pit at E10.5. Summary: The αA-crystallin ranks amongst the most highly expressed tissue-specific genes. It is an advantageous model system to probe both promoter-enhancer looping and to identify distal enhancers and their temporal/spatial activities. AlthougAn advantageous tissue to study transcription, chromatin dynamics and subnuclear organization and compartmentalization during mammalian development is the ocular lens. The mature lens consists of an anterior layer of epithelial cells that overlies the bulk of the lens, made up of differentiated fiber cells. This compartmentalization of the lens into epithelium and fibers initiates from an early transitional structure termed the lens vesicle . DiffereCryaa) gene evolved from gene duplication of the αB-crystallin (Cryab) that evolved from Hspb1-like ancestral gene   and a paPrevious studies of the temporal regulation of αA-crystallin did not identify the DNA region necessary for its earliest expression in the lens pit . Exogenon=3). It was observed that a significant interaction occurs between the promoter and a large portion of the chromatin 3′ adjacent to DCR1 (n=2) and found a similar broadened peak at the DCR1 region as well as the 3′ distal peak . We extended the region examined by 10 kb 3′ beyond this distal peak and observed no further interaction peaks (data not shown). Based on this data we hypothesized that the region 3′ to DCR1  could be important for regulation of αA-crystallin expression, possibly at the earlier stages of E10.5.Recent work has demonstrated the importance of chromatin conformation on gene expression with significant progress being made in determining looping that occurs within gene loci to allow for interactions between enhancer and promoter regions required for gene transcription . To dete to DCR1 B. This i to DCR1 C as wellTo determine if there are any differences between two lens cell types, we next examined looping in chromatin from microdissected newborn (P1) lens epithelium and fiber cells . As withPrior analysis of mice expressing EGFP under control of the αA-crystallin 1.9 kb genomic region containing the promoter and its 5′-adjacent DCR2 show modest expression in lens fiber cells from E13.5 . To deteIn this study we present evidence for DNA looping at the αA-crystallin locus in mouse. Based on this chromatin conformation we subsequently identified a novel enhancer region, which, determined by transgenic mouse studies, is thought to act as a shadow enhancer eliciting expression patterns similar to that of DCR1. Although expression of αA-crystallin is highly tissue-restricted, looping was also found in terminally differentiated non-αA-crystallin expressing tissues, albeit at a lower frequency. In contrast, the pluripotent ES cells do not exhibit any significant physical interactions of distal regions within the αA-crystallin locus.polyA, DCR3 absent; Due to the short window of time in which it has to produce all proteins required for the life of the organism, lens is an advantageous model tissue in which to study transcriptional control. αA-crystallin in particular, with its extraordinarily high relative expression levels must be highly regulated to generate such massive quantities before fiber cell organelles are degraded . PreviouS1), suggesting the existence of a novel 1.5 kb regulatory region, from here on referred to as ‘DCR1S’. Interaction frequency between the promoter and DCR1S was much higher at the later developmental stages of E15.5 and P1 compared to E14.5  (snail (The phenomenon of two or more enhancer regions that are able to perform similar functions is known as ‘enhancer redundancy’ . Studiesosophila . The evoby (svb)  and snai) (snail  genes ap) (snail  or tempo)  in non-lens tissues. This suggests the presence of an enhancer RNA (eRNA) transcribed from DCR1. However, our attempts to detect this transcript in mouse tissues based on RNA-seq data and RT-PCR did not reveal the presence of an eRNA (data not shown). Evolutionary differences acting on DCR1 and DCR1S enhancers in mouse and human are likely to explain presence or absence of eRNAs and can be further experimentally tested, including studies of nd DCR1S  and examnd DCR1S .Cryaa locus. These enhancers are both capable of inducing expression in the lens vesicle at mouse stage E11.5, however it remains to be elucidated which region of the αA-crystallin locus is responsible for expression in the lens pit at E10.5. Possible candidates include the 3′ distal region for which we see looping with the promoter as well as other evolutionarily conserved noncoding regions identified in introns 1 and 2 . The 15 kb band was excised from the gel and melted in TAE buffer. The DNA was precipitated in 100% ethanol and sodium acetate , then washed with 70% ethanol. The 15 kb fragment was cloned into pBluescript SKII vector sites, XmaI and SpeI. 15 kb αA-ΔDCR1 ΔDCR3-GFP construct . Embryos were genotyped by PCR using primers against EGFP . Three lines of each mouse were analyzed and demonstrated similar expression patterns  as diagrammatically shown in patterns .® 488 (1:500) (Molecular Probes). Slides were washed and mounted with Vectashield containing DAPI (Vector). Images were taken with a Leica AOBS laser scanning confocal microscope.Embryos were fixed in 4% paraformaldehyde, cryoprotected with 15% sucrose in PBS, and embedded in Optimal Cutting Temperature (OCT) tissue freezing medium . Transverse cryostat sections (10 μm) were collected, washed with PBS containing 0.1% triton-X, blocked for 1 h in PBS containing 1% BSA, and then incubated overnight at 4°C with the primary antibody, rabbit anti-GFP , diluted in PBS containing 1% BSA. Sections were washed twice for 10 min in PBS containing 0.1% triton-X and incubated for 45 min with the secondary antibody, goat anti-rabbit Alexa FluorRNA was isolated from dissected tissues using miRNesay mini kit (Qiagen) according to manufacturer's instructions. cDNA was generated with Superscript™ III Reverse Transcriptase (Thermo Fisher), and the template was diluted 1:10. Quantitative PCR was performed using SYBR Green (Thermo Fisher) and primers: αA-crystallin (5′-GAGATTCACGGCAAACACAA-3′ and 5′-ACATTGGAAGGCAGACGGTA-3′) and EGFP . Primers for αA-crystallin recognize both endogenous and the fusion αA-crystallin-EGFP cDNA. The relative expression level of αA-crystallin was normalized by the EGFP fusion protein average versus endogenous αA-crystallin. Three lines of each mouse were analyzed and EGFP expression values can be seen in Genomic DNA was isolated by digesting tissue with lysis buffer  containing Proteinase K (100 μg/ml) at 55°C. Phenol/chloroform/isoamyl alcohol (Thermo Fisher) extractions were performed and DNA was precipitated with isopropanol. Quantitative PCR was conducted using SYBR Green (Thermo Fisher) to determine the number of copies of the transgene, using the following primers: αA-crystallin (5′-GAGAGGGCCATTCCTGTGT-3′ and 5′-AGGGGACAACCAAGGTGAG-3′); (5′-GGGTGCTGGTCTACTTCCAG-3′ and 5′-AACCACGACATCCGAAAAAG-3′); and CCNI (5′-TCTTCTCCCTCCTCAGACG-3′ and 5′-CCGTTACCACCTCATGATCC-3′); B2M (5′-CCCTGGCTGGCTCTCATT-3′ and 5′-ACTGAAGCGACCGCGACT-3′) for normalization. Three lines of each mouse were analyzed and copy number ranges can be seen in 2O and 100 μl 20% Triron X-100  was added and incubated at 37°C for 1 h with shaking. The reaction mixture was then cooled to 16°C and 50 U of T4 DNA ligase (NEB) was added. After 4 h of ligation, the chromatin mixture was incubated with 100 μg/ml proteinase K at 65°C overnight to reverse cross-links. RNA was removed by RNase A (0.5 μg/ml) treatment for 30 min at 37°C and DNA was purified by phenol extraction. Quantitative real-time PCRs were performed, in the presence of SYBR Green (Thermo Fisher), with appropriate primers (sequences available upon request) from purified DNA as well as the control template. The control template was generated by digestion and ligation of αA-Crystallin BAC clone (RP-23-465G4) to generate each ligation product in equal molar amounts. The relative cross-linking frequency between two fragments was calculated by normalizing to the control library. This calculation corrects for differences in cross-linking and ligation efficiencies, PCR amplification efficiency, the amount of the initial template used and the sizes of the PCR products.3C assays were carried out as previously described , with soRNA was isolated from P1 mouse lens, liver, forebrain, heart and mouse embryonic stem (mES) cells using miRNesay mini kit (Qiagen) according to manufacturer's instructions. cDNA was generated with Superscript™ III Reverse Transcriptase (Thermo Fisher). Quantitative PCR using SYBR Green (Thermo Fisher) was performed to determine the levels of αA-crystallin and normalized to Gapdh and B2m using the following primers: αA-crystallin (Cryaa: 5′-GAGATTCACGGCAAACACAA-3′ and 5′-ACATTGGAAGGCAGACGGTA-3′); Gapdh ; B2m .