The MerR-family transcription factors (TFs) are a large group of bacterial proteins responding to cellular metal ions and multiple antibiotics by binding within central RNA polymerase-binding regions of a promoter. While most TFs alter transcription through protein–protein interactions, MerR TFs are capable of reshaping promoter DNA. To address the question of which mechanism prevails, we determined two cryo-EM structures of transcription activation complexes (TAC) comprising Escherichia coli CueR (a prototype MerR TF), RNAP holoenzyme and promoter DNA. The structures reveal that this TF promotes productive promoter–polymerase association without canonical protein–protein contacts seen between other activator proteins and RNAP. Instead, CueR realigns the key promoter elements in the transcription activation complex by clamp-like protein–DNA interactions: these induce four distinct kinks that ultimately position the −10 element for formation of the transcription bubble. These structural and biochemical results provide strong support for the DNA distortion paradigm of allosteric transcriptional control by MerR TFs.
Subscribe to Journal
Get full journal access for 1 year
only $14.08 per issue
All prices are NET prices.
VAT will be added later in the checkout.
Rent or Buy article
Get time limited or full article access on ReadCube.
All prices are NET prices.
Browning, D. F., Butala, M. & Busby, S. J. W. Bacterial transcription factors: regulation by pick ‘n’ mix. J. Mol. Biol. 431, 4067–4077 (2019).
Danson, A. E., Jovanovic, M., Buck, M. & Zhang, X. Mechanisms of σ54-dependent transcription initiation and regulation. J. Mol. Biol. 431, 3960–3974 (2019).
Feng, Y., Zhang, Y. & Ebright, R. H. Structural basis of transcription activation. Science 352, 1330–1333 (2016).
Liu, B., Hong, C., Huang, R. K., Yu, Z. & Steitz, T. A. Structural basis of bacterial transcription activation. Science 358, 947–951 (2017).
Hubin, E. A. et al. Structure and function of the mycobacterial transcription initiation complex with the essential regulator RbpA. eLife 6, e22520 (2017).
Bae, B. et al. CarD uses a minor groove wedge mechanism to stabilize the RNA polymerase open promoter complex. eLife 4, e08505 (2015).
Brown, N. L., Stoyanov, J. V., Kidd, S. P. & Hobman, J. L. The MerR family of transcriptional regulators. FEMS Microbiol. Rev. 27, 145–163 (2003).
Fernandez-Lopez, R., Ruiz, R., de la Cruz, F. & Moncalian, G. Transcription factor-based biosensors enlightened by the analyte. Front. Microbiol. 6, 648 (2015).
Kulkarni, R. D. & Summers, A. O. MerR cross-links to the σ, β, and σ70 subunits of RNA polymerase in the preinitiation complex at the merTPCAD promoter. Biochemistry 38, 3362–3368 (1999).
Ishihama, A. in Nucleic Acids and Molecular Biology, Vol. 11 (eds Eckstein, F. & Lilley D. M. J.) 53–70 (Springer-Verlag, 1997).
Heldwein, E. E. & Brennan, R. G. Crystal structure of the transcription activator BmrR bound to DNA and a drug. Nature 409, 378–382 (2001).
Philips, S. J. et al. Allosteric transcriptional regulation via changes in the overall topology of the core promoter. Science 349, 877–881 (2015).
Ansari, A. Z., Bradner, J. E. & O’Halloran, T. V. DNA-bend modulation in a repressor-to-activator switching mechanism. Nature 374, 371–375 (1995).
Ansari, A. Z., Chael, M. L. & O’Halloran, T. V. Allosteric underwinding of DNA is a critical step in positive control of transcription by Hg-MerR. Nature 355, 87–89 (1992).
Watanabe, S., Kita, A., Kobayashi, K. & Miki, K. Crystal structure of the [2Fe-2S] oxidative-stress sensor SoxR bound to DNA. Proc. Natl Acad. Sci. USA 105, 4121–4126 (2008).
Outten, C. E., Outten, F. W. & O’Halloran, T. V. DNA distortion mechanism for transcriptional activation by ZntR, a Zn(II)-responsive MerR homologue in Escherichia coli. J. Biol. Chem. 274, 37517–37524 (1999).
O’Halloran, T. V., Frantz, B., Shin, M. K., Ralston, D. M. & Wright, J. G. The MerR heavy metal receptor mediates positive activation in a topologically novel transcription complex. Cell 56, 119–129 (1989).
Frantz, B. & O’Halloran, T. V. DNA distortion accompanies transcriptional activation by the metal-responsive gene-regulatory protein MerR. Biochemistry 29, 4747–4751 (1990).
Martell, D. J. et al. Metalloregulator CueR biases RNA polymerase’s kinetic sampling of dead-end or open complex to repress or activate transcription. Proc. Natl Acad. Sci. USA 112, 13467–13472 (2015).
Zhang, Y. et al. Structural basis of transcription initiation. Science 338, 1076–1080 (2012).
Bae, B., Feklistov, A., Lass-Napiorkowska, A., Landick, R. & Darst, S. A. Structure of a bacterial RNA polymerase holoenzyme open promoter complex. eLife 4, e08504 (2015).
Narayanan, A. et al. Cryo-EM structure of Escherichia coli σ70 RNA polymerase and promoter DNA complex revealed a role of σ non-conserved region during the open complex formation. J. Biol. Chem. 293, 7367–7375 (2018).
Shultzaberger, R. K., Chen, Z., Lewis, K. A. & Schneider, T. D. Anatomy of Escherichia coli σ70 promoters. Nucleic Acids Res. 35, 771–788 (2007).
Campbell, E. A. et al. Structure of the bacterial RNA polymerase promoter specificity σ subunit. Mol. Cell 9, 527–539 (2002).
Mekler, V., Pavlova, O. & Severinov, K. Interaction of Escherichia coli RNA polymerase σ70 subunit with promoter elements in the context of free σ70, RNA polymerase holoenzyme, and the β’-σ70 complex. J. Biol. Chem. 286, 270–279 (2011).
Mekler, V. & Severinov, K. RNA polymerase molecular beacon as tool for studies of RNA polymerase–promoter interactions. Methods 86, 19–26 (2015).
Hudson, B. P. et al. Three-dimensional EM structure of an intact activator-dependent transcription initiation complex. Proc. Natl Acad. Sci. USA 106, 19830–19835 (2009).
Rammohan, J., Ruiz Manzano, A., Garner, A. L., Stallings, C. L. & Galburt, E. A. CarD stabilizes mycobacterial open complexes via a two-tiered kinetic mechanism. Nucleic Acids Res. 43, 3272–3285 (2015).
Rammohan, J. et al. Cooperative stabilization of Mycobacterium tuberculosis rrnAP3 promoter open complexes by RbpA and CarD. Nucleic Acids Res. 44, 7304–7313 (2016).
Jensen, D., Manzano, A. R., Rammohan, J., Stallings, C. L. & Galburt, E. A. CarD and RbpA modify the kinetics of initial transcription and slow promoter escape of the Mycobacterium tuberculosis RNA polymerase. Nucleic Acids Res. 47, 6685–6698 (2019).
Ralston, D. M. & O’Halloran, T. V. Ultrasensitivity and heavy-metal selectivity of the allosterically modulated MerR transcription complex. Proc. Natl Acad. Sci. USA 87, 3846–3850 (1990).
O’Halloran, T. V. Transition metals in control of gene expression. Science 261, 715–725 (1993).
Outten, F. W., Outten, C. E., Hale, J. & O’Halloran, T. V. Transcriptional activation of an Escherichia coli copper efflux regulon by the chromosomal MerR homologue, cueR. J. Biol. Chem. 275, 31024–31029 (2000).
Gaston, K., Bell, A., Kolb, A., Buc, H. & Busby, S. Stringent spacing requirements for transcription activation by CRP. Cell 62, 733–743 (1990).
Fang, C. et al. Structures and mechanism of transcription initiation by bacterial ECF factors. Nucleic Acids Res. 47, 7094–7104 (2019).
Li, L., Fang, C., Zhuang, N., Wang, T. & Zhang, Y. Structural basis for transcription initiation by bacterial ECF σ factors. Nat. Commun. 10, 1153 (2019).
Zheng, S. Q. et al. MotionCor2: anisotropic correction of beam-induced motion for improved cryo-electron microscopy. Nat. Methods 14, 331–332 (2017).
Zhang, K. Gctf: real-time CTF determination and correction. J. Struct. Biol. 193, 1–12 (2016).
Fernandez-Leiro, R. & Scheres, S. H. W. A pipeline approach to single-particle processing in RELION. Acta Crystallogr. D Struct. Biol. 73, 496–502 (2017).
Degen, D. et al. Transcription inhibition by the depsipeptide antibiotic salinamide A. eLife 3, e02451 (2014).
Henderson, R. et al. Outcome of the first electron microscopy validation task force meeting. Structure 20, 205–214 (2012).
Emsley, P. & Cowtan, K. Coot: model-building tools for molecular graphics. Acta Crystallogr. D Biol. Crystallogr. 60, 2126–2132 (2004).
Adams, P. D. et al. PHENIX: a comprehensive Python-based system for macromolecular structure solution. Acta Crystallogr. D Biol. Crystallogr. 66, 213–221 (2010).
Lavery, R., Moakher, M., Maddocks, J. H., Petkeviciute, D. & Zakrzewska, K. Conformational analysis of nucleic acids revisited: Curves+. Nucleic Acids Res. 37, 5917–5929 (2009).
Lu, X. J. & Olson, W. K. 3DNA: a versatile, integrated software system for the analysis, rebuilding and visualization of three-dimensional nucleic-acid structures. Nat. Protoc. 3, 1213–1227 (2008).
Pettersen, E. F. et al. UCSF Chimera—a visualization system for exploratory research and analysis. J. Comput. Chem. 25, 1605–1612 (2004).
Murakami, K. S., Masuda, S. & Darst, S. A. Structural basis of transcription initiation: RNA polymerase holoenzyme at 4 Å resolution. Science 296, 1280–1284 (2002).
Boyaci, H., Chen, J., Jansen, R., Darst, S. A. & Campbell, E. A. Structures of an RNA polymerase promoter melting intermediate elucidate DNA unwinding. Nature 565, 382–385 (2019).
Feklistov, A. et al. RNA polymerase motions during promoter melting. Science 356, 863–866 (2017).
Boyaci, H. et al. Fidaxomicin jams Mycobacterium tuberculosis RNA polymerase motions needed for initiation via RbpA contacts. eLife 7, e34823 (2018).
Lin, W. et al. Structural basis of transcription inhibition by fidaxomicin (lipiarmycin A3). Mol. Cell 70, 60–71.e15 (2018).
Chakraborty, A. et al. Opening and closing of the bacterial RNA polymerase clamp. Science 337, 591–595 (2012).
Trachman, R. J. III et al. Structural basis for high-affinity fluorophore binding and activation by RNA Mango. Nat. Chem. Biol. 13, 807–813 (2017).
Miller, J. H. Experiments in Molecular Genetics (Cold Spring Harbor Laboratory, 1972).
The work was supported by the National Key Research and Development Program of China (grant no. 2018YFA0900701 to Y.Z. and grant no. 2018YFA0507800 to Y.F.), the Strategic Priority Research Program of CAS to Y.Z. (grant no. XDB29020000), the National Natural Science Foundation of China (grant no. 31822001 to Y.Z. and grant no. 31970040 to Y.F.), the Leading Science Key Research Program of CAS to Y.Z. (grant no. QYZDB-SSW-SMC005) and the National Institutes of Health of the United States to T.V.O. (grant nos. GM038784-31 and CA193419). We thank S. Chang at the cryo-EM center of Zhejiang University, L. Kong and F. Wang at the cryo-EM center of the National Center for Protein Science Shanghai for assistance with data collection, C. Wang at Rutgers University for assistance with figure preparation and A. Mondragón at Northwestern University for helpful discussions.
The authors declare no competing interests.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
(a) template strand (TS); (b) non-template strand (NTS). For DNase I footprinting, the concentrations of CueR, Cu(I) or Ag(I) [M(I)], CN- and RNAP were 50 nM, 50 μM, 1 mM and 50 nM, respectively. For KMnO4 footprinting, the concentrations of CueR, M(I), TPEN and RNAP were 50 nM, 5 μM, 100 μM and 5 nM, respectively. For 5-phenyl-1,10-phenanthroline-Cu footprinting, the concentrations of CueR, TPEN and RNAP were 50 nM, 10 μM and 5 nM, The metal-free complex was not interrogated by 5-phenyl-1,10-phenanthroline-Cu footprinting due to the inducing ion, Cu, being part of the probe. In lanes 8, 16, 17, 18 and 22, 100 μM of dinucleotide ApG and 10 μM of ATP, CTP and GTP nucleotides were added. Lanes G are guanines-specific sequence ladders. The DNaseI footprint is reconciled with the cryo-EM structures, both showing overlapping coverage of the promoter DNA by CueR and RNAP. The KMnO4 probe is used to identify unpaired thymidine bases (T). Several aspects KMnO4 footprint also match the structural nature of the CueR-TAC-1 structure, specifically in the transcription bubble. Our analysis focuses on CueR-TAC-1 due to the higher resolution and the presence of the template and non-template strands in the transcription bubble. TS −2T (G in the DNA used for EM); the upstream half of TS −10 element (ATT; AAT in the cryo-EM DNA); the downstream area of TS −10 element TGG in the cryo-EM DNA): these are all hypersensitive to KMnO4 in the footprint, and are observed to be unstacked in the structure, and thus subject to KMnO4 attack. In contrast, the template strand T at +1 (also T in the cryo-EM DNA) is not hypersensitive to KMnO4 in the footprint even though it is in the transcription bubble. This agrees with the structure, which shows clear stacking interactions with the adjacent bases. The representative gel image was from one biologically independent experiment. Source data
a, The nucleic-acid scaffold used for preparing the CueR-TAC-1 in the study. The CueR operator DNA is highlighted in a dashed box and the palindromic sequence is underlined by green arrows. The nontemplate ssDNA, orange; the template ssDNA, red; the −35 element, light blue; the −10 element, violet; the discriminator element; cyan; the core-recognition element, green; RNA, blue. b, The elution peaks of E. coli CueR-TAC from a size-exclusion column. c, The SDS-PAGE of the E. coli CueR-TAC-1 complex, CueR, and σ70. PAGE of the E. coli CueR-TAC-1 complex stained with Coomassie Brilliant Blue or SYBR Gold dyes. The SDS-PAGE and native-PAGE images were from a single experiment. e, The in vitro transcription activity showing the E. coli CueR activates transcription from PcopA in the presence of AgNO3. The gel image was from a single experiment. f, The PcopA derivative used for cryo-EM structure determination of E. coli CueR-TAC-2. The PcopA derivative comprises wild-type sequence of PcopA (−60 to −11) and consensus sequences for the rest of the −10, discriminator, and core-recognition elements (gray-shaded). Colors as in (a). g, RPo formation from the PcopA derivative (f) in a gel-shift assay. The asterisk labels aggregated protein. RPc, RNAP-promoter DNA closed complex. The representative gel image was from one of three biologically independent experiments. h, CueR facilitates RPo formation from the PcopA derivative (f) in a molecular beacon assay. Data were plotted as mean ± SEM. n= 3 biologically independent samples. (i) CueR activates transcription from the PcopA derivative in a fluorescence-detected in vitro transcription assay. Data were plotted as mean ± SEM. n= 5 biologically independent samples. Source data
a, The superimposition of CueR-DNA of the two structures. The RMSD value of the CueR Cα atoms of two structures is 2.0 Å. The E. coli CueR-TAC-2 is colored in light green; the CueR dimer of E. coli CueR-TAC-1 is colored in green and cyan; and the DNA of E. coli CueR-TAC-1 is colored in red and orange. b, The superimposition of RNAP holoenzyme of the two structures. The RMSD values of all RNAP Cα atoms of the two structures is 3.0 Å. The E. coli CueR-TAC-2 is colored in light green and the RNAP-β’, β, σ of E. coli CueR-TAC-1 are colored in black, gray, and blue respectively. c, The DNA conformation is essentially identical of the two structures except that the CueR-TAC-2 shows ordered “UP” element and disordered template ssDNA in the transcription bubble. The E. coli CueR-TAC-2 is colored in light green. The non-template DNA, template DNA, RNA strands of E. coli CueR-TAC-1 are colored in orange, red, and blue respectively. d, The αCTD of E. coli CueR-TAC-1 makes interaction with both the UP element and σ4 but makes no interaction with CueR. The colors are as in Fig. 2e.
Extended Data Fig. 4 The interactions of σ704 and αCTD with the −35 and UP elements, respectively, in the CueR-TAC-2 structure are physiologically important.
a, Two view-orientations of cartoon representations of the interactions of σR4 (blue) with the −35 promoter element and of αCTD (gold) with the UP element. b, Curves+ analysis of the major (solid lines) and minor grooves (dashed lines) in the upstream region of the CueR-TAC-2 structure, encompassing UP elements; and, comparison to the groove widths in idealized B-DNA. The UP element region (TAATTT) exhibits clear reduced minor groove width relative to ideal B-DNA, which reduces the likelihood of DNaseI digestion. This in agreement with the DNaseI footprinting pattern seen in Fig. 1 and Extended Data Fig. 1, as the UP element is nearly fully protected from DNase I digestion. Moreover, the first half of the −35 promoter element hexamer (TTG) exhibits increased minor groove width relative to ideal B-DNA. Finally, the second half of the −35 promoter element hexamer (ACC) exhibits decreased minor groove width relative to ideal B-DNA, and is fully covered by the wing of the DNA-binding domain of the upstream CueR protomer. DNAase I has higher activity at sites with wider groove widths; this analysis of the structure predicts DNase I sensitivity at these base steps, and the corresponding DNase I footprinting assays reveal hypersensitivity at positions −40A of the template strand, and −36G and −37T of the non-template strand (Fig. 1 and Extended Data Fig. 1). We conclude that key structural features of the observed in the cryo-EM sample are also present in CueR-complexes in physiologically relevant buffers at room temperature.
a, CueR DBD α2 residues K15 and F19 insert into the major groove and interact with DNA bases at the center of the operator. b, CueR wing loop residue Y36 inserts into the minor groove and interacts with −35 element DNA bases at the distal edge of the operator. c, Several CueR DBD residues also make extensive interactions with DNA backbone phosphates, stabilizing the protein-DNA interaction. d, The interaction between σ704 and the consensus sequence “TTG” of the −35 element. e, The structure superimposition of T. aquaticus σ704/−35 binary structure (brown; PDB: 1KU7) and E. coli CueR-TAC (red and blue). f, The interaction between the extended −10 region of promoter DNA and σ70. The cryo-EM map for sidechains of amino acids is shown as blue mesh. The nontemplate ssDNA, orange; the template ssDNA, red; CueRU-DBD, cyan; CueRD-DBD, green. σ70, blue; green dashed lines, H-bonds.
The sequence logo was generated by WebLogo V2.8.2 (https://weblogo.berkeley.edu).
a, Cartoon representation of the activator CueR dimer (CueRU and CueRD protomers) bound to promoter DNA in CueR-TAC-1. (RNAP holoenzyme has been removed for clarity). Activator CueR binds to its cognate operator and kinks the DNA at the center of the promoter. The various elements are colored and labeled as in Fig. 2. The starred base pairs in cartoon correspond to the starred base pairs in B. b, The kinks at the center of the upstream dsDNA in the E. coli CueR-TAC-1 and CueR-TAC-2 structures are very similar to the kink observed in the activator CueR/DNA crystal structure (PDB: 4WLW) in (c). The overall kinks are represented by the roll angles over the two central base pair steps [T:A – T:A – G:C], which add up to ~53° in the E. coli CueR-TAC-1 and ~52° in CueR-TAC-2 structure (vs. ~55° in the activator CueR/DNA structure). d, Major (M) and (m) groove widths of DNA in the copA operator in the CueR-TAC-1 (blue), CueR-TAC-2 (purple) and activator CueR/DNA structures (green). The major and minor groove widths are very similar in both structures, but both groove widths increase near the −10 element site (labeled and arrowed above the DNA sequence), as expected due to the “pull” exerted on the DNA by the formation of the transcription bubble by the RNA polymerase. e, Overlays (using the UCSF Chimera X matchmaker tool) of CueR-TAC-1 and the CueR/DNA/RNAP complex model (left)12, and CueR-TAC-2 and the Activator CueR/DNA/RNAP complex model12. The overall RMSD of the model vs. CueR-TAC-1 and vs. TAC-2 are 8.45Å and 8.2Å across all atoms, respectively. The largest differences between the modeled CueR/DNA/RNAP complex and the two TAC structure are DNA beginning ca. 6 bp upstream of the transcription bubble. While no melted DNA was introduced in the model, the duplex promoter DNA downstream of kink is pulled down and melted in the CueR-TAC-1 and CueR-TAC-2 structures. The cryo-EM structures presented herein strongly validate the predictions for the Activator CueR/DNA/RNAP complex model made previously12.
a, No interaction between CueRU and σ704. b, The potential interaction between CueRD and σ70NCR. c, Results of the β-gal assay for variant E33A, which shows wild-type transcriptional response to all added CuSO4 concentrations. Data were plotted as mean ± SEM. n= 6 biologically independent samples except for the variant E33A (n=3). d, No interaction between the CueR dimer and RNAP core enzyme.
a, The sequence alignment of the MerR family TFs. b, The structure modeling of B. subtilis TAC. The crystal structure of B. subtilis BmrR/dsDNA (PDB:3Q2Y) was superimposed on our cryo-EM structure of E. coli CueR-TAC. The structure superimposition shows that the large C-terminal multidrug binding domain extrudes out and makes no interactions with RNAP holoenzyme. Yellow and orange surfaces, solvent-accessible surfaces of RNAP-α subunits; gray, dark gray, pink, and blue surfaces, solvent-accessible surfaces of RNAP-β, RNAP-β′, RNAP-ω subunit, and σA, respectively. Orange ribbon, promoter DNA; green and cyan ribbons, BmrR dimer.
About this article
Cite this article
Fang, C., Philips, S.J., Wu, X. et al. CueR activates transcription through a DNA distortion mechanism. Nat Chem Biol (2020). https://doi.org/10.1038/s41589-020-00653-x