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Glycoproteomics is coming of age, thanks to advances in instrumentation, experimental methodologies and computational search algorithms.

Glycosylation is one of the most common post-translational modifications, and glycoproteins play crucial roles in important biological processes like cell signaling, host–pathogen interaction, immune response and disease, including cancer and even the ongoing COVID-19 pandemic (Science 369, 330–333, 2020). Glycoproteomics aims to determine the positions and identities of the complete repertoire of glycans and glycosylated proteins in a given cell or tissue.

Glycans are everywhere. High-throughput glycoproteomics approaches offer insights. Credit: Katherine Vicari, Springer Nature

Mass spectrometry (MS)-based approaches allow large-scale global analysis; however, the structural diversity of glycans and the heterogeneous nature of glycosylation sites make comprehensive analysis particularly challenging. Glycans obstruct complete fragmentation of the protein backbone, and they were traditionally removed for simplicity at the cost of losing glycan information. The MS spectra tend to be complicated due to the presence of isomers, often requiring manual interpretation. Furthermore, database searching for spectral matches can quickly become a combinatorial problem and requires innovative bioinformatics solutions.

Recent developments in MS instrumentation, fragmentation strategies (J. Proteome Res. 19, 3286–3301, 2020) and high-throughput workflows have made analyzing intact glycoproteins a possibility. Several specific enrichment strategies have made even low-abundance glycans and glycopeptides detectable (Mol. Cell. Proteomics, 2020). A variety of experimental workflows tailored for either N-linked glycans, which are found at consensus sites on the proteins, or O-linked glycans, which have no recognizable consensus sequence, have been developed (Nature 549, 538–542, 2017; Nat. Commun. 11, 5268, 2020; Nat. Methods 16, 902–910, 2019). New software packages based on fragment-ion indexing strategies offer substantial increases in speed for glycopeptide and site assignments (Nat. Methods 17, 1125–1132, 2020; Nat. Methods 17, 1133–1138, 2020).

With other -omics fields taking the lion’s share of attention in recent years, it is now time for glycoproteomics to shine. Comprehensive understanding of glycosylation at different levels of granularity is bound to serve both basic and translational research.

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Correspondence to Arunima Singh.

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Singh, A. Glycoproteomics. Nat Methods 18, 28 (2021).

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Bio Chemistry

Structures of the glucocorticoid-bound adhesion receptor GPR97–Go complex

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No statistical methods were used to predetermine sample size. The experiments were not randomized, and investigators were not blinded to allocation during experiments and outcome assessment.

Cell lines

HEK293 cells were obtained from the Cell Resource Center of Shanghai Institute for Biological Sciences (Chinese Academy of Sciences). Spodoptera frugiperda (Sf9) cells were purchased from Expression Systems (cat. 94-001S). Y-1 cells were originally obtained from the American Type Culture Collection (ATCC). The cells were grown in monolayer culture in RPMI 1640 with 10% FBS (Gibco) at 37 °C in a humidified atmosphere consisting of 5% CO2 and 95% air.

Constructs of GPR97 and miniGo heterotrimer

For protein production in insect cells, the human GPR97 (residues 21–549) with the autoproteolysis motif mutation (H248/A and T250/A) was sub-cloned into the pFastBac1 vector. The native signal peptide was replaced with the haemagglutinin signal peptide (HA) to enhance receptor expression, followed by a Flag tag DYKDDDK (China peptide) to facilitate complex purification. An engineered human Gαo1 with Gαo1 H domain deletion, named miniGαo1 was cloned into pFastBac1 according to published literature29. Human Gβ1 with the C-terminal hexa-histidine tag and human Gγ2 were subcloned into the pFastBacDual vector. scFv16 was cloned into pfastBac1 with the C-terminal hexa-histidine tag and the N-terminal GP67 signal peptide. To examine the activities of GPR97, the GPR97-FL-WT (wild-type full-length GPR97), GPR97-FL-AA (GPR97 GPS site mutation, H248/A and T250/A), GPR97β (GPR97 with the NTF removed, residues 250–549) and GPR97-β-T (GPR97β with the N-terminal tethered Stachel sequence removed, residues 265–549) were sub-cloned into the pcDNA3.1 plasmid. The GPR97 mutations E298A, R299A, F345A, F353A, H362A, L363A, Y364A, V370A, F371A, Y406A, W421A, W490A, A493G, I494A, L498A and N510A were generated using the Quikchange mutagenesis kit (Stratagene). The G protein BRET probes were constructed according to previous publications42,43. Human G protein subunits (Gαq, Gβ1 and Gγ2) were sub-cloned into the pcDNA3.1 expression vectors. The Gαq-RlucII subunit was generated by amplifying and inserting the coding sequence of RlucII into Gαq between residue L97 and K98. The Gqo probe, in which the six amino acids of the C-terminal of Gαq-RlucII were substituted with those from Gαo1, was constructed by PCR amplification using synthesized oligonucleotides encoding swapped C-terminal sequences. The GFP10–Gγ2 plasmid was generated by fusing the GFP10 coding sequence in frame at the N terminus to Gγ2. All of the constructs and mutations were verified by DNA sequencing.

Protein expression

High titre recombinant baculoviruses were generated using Bac-to-Bac Baculovirus Expression System. In brief, 2 μg of recombinant bacmid and 2 μl X-tremGENE HP transfection reagent (Roche) in 100 μl Opti-MEM medium (Gibco) were mixed and incubated for 20 min at room temperature. The transfection solution was added to 2.5 ml Sf9 cells with a density of 1 × 106 per ml in a 24-well plate. The infected cells were cultured in a shaker at 27 °C for 4 days. P0 virus was collected and then amplified to generate P1 virus. The viral titres were determined by flow cytometric analysis of cells stained with gp64-PE antibody (1:200 dilution; 12-6991-82, Thermo Fisher). Then, Sf9 cells were infected with viruses encoding GPR97-FL-AA, miniGαo, Gβγ, and with or without scFv16, respectively, at equal multiplicity of infection. The infected cells were cultured at 27 °C, 110 rpm for 48 h before collection. Cells were finally collected by centrifugation and the cell pellets were stored at −80 °C.

GPR97–Go complex formation and purification

Cell pellets transfected with virus encompassing the GPR97-FL-AA, miniGo trimer and scFv16 (only existed in cell pellets for purifying the cortisol–GPR97-FL-AA–Go–scFv16 complex) were resuspended in 20 mM HEPES, pH 7.4, 100 mM NaCl, 10% glycerol, 10 mM MgCl2 and 5 mM CaCl2 supplemented with Protease Inhibitor Cocktail (B14001, Bimake) and 100 μM TCEP (Thermo Fisher Scientific). The complex was formed for 2 h at room temperature by adding 10 μM BCM (HY-B1540, MedChemExpress) or cortisol (HY-N0583, MedChemExpress), 25 mU/ml apyrase (Sigma), and then solubilized by 0.5% (w/v) lauryl maltose neopentylglycol (LMNG; Anatrace) and 0.1% (w/v) cholesteryl hemisuccinate TRIS salt (CHS; Anatrace) for 2 h at 4 °C. Supernatant was collected by centrifugation at 30,000 rpm for 40 min, and the solubilized complex was incubated with nickel resin for 2 h at 4 °C. The resin was collected and washed with 20 column volumes of 20 mM HEPES, pH 7.4, 100 mM NaCl, 10% glycerol, 2 mM MgCl2, 25 mM imidazole, 0.01% (w/v) LMNG, 0.01% GDN (Anatrace), 0.004% (w/v) CHS, 10 μM BCM (or cortisol) and 100 μM TCEP. The complex was eluted with 20 mM HEPES, pH 7.4, 100 mM NaCl, 10% glycerol, 2 mM MgCl2, 200 mM imidazole, 0.01% (w/v) LMNG, 0.01% GDN, 0.004% (w/v) CHS, 10 μM BCM (or cortisol) and 100 μM TCEP. The elution of nickel resin was applied to M1 anti-Flag resin (Sigma) for 2 h and washed with 20 mM HEPES, pH 7.4, 100 mM NaCl, 10% glycerol, 2 mM MgCl2, 5 mM CaCl2, 0.01% (w/v) LMNG, 0.01% GDN, 0.004% (w/v) CHS, 10 μM BCM (or cortisol) and 100 μM TCEP. The GPR97–Go complex was eluted in buffer containing 20 mM HEPES, pH 7.4, 100 mM NaCl, 10% glycerol, 2 mM MgCl2, 0.01% (w/v) LMNG, 0.01% GDN, 0.004% (w/v) CHS, 10 μM BCM (or cortisol), 100 μM TCEP, 5 mM EGTA and 0.2 mg/ml Flag peptide. The complex was concentrated and then injected onto Superdex 200 increase 10/300 GL column equilibrated in the buffer containing 20 mM HEPES, pH 7.4, 100 mM NaCl, 2 mM MgCl2, 0.00075% (w/v) LMNG, 0.00025% GDN, 0.0002% (w/v) CHS, 10 μM BCM (or cortisol) and 100 μM TCEP. The complex fractions were collected and concentrated individually for EM experiments.

Cryo-EM grid preparation and data collection

For the preparation of cryo-EM grids, 3 μl of purified BCM-bound and cortisol-bound GPR97–Go complex at approximately 20 mg/ml was applied onto a glow-discharged holey carbon grid (Quantifoil R1.2/1.3). Grids were plunge-frozen in liquid ethane cooled by liquid nitrogen using Vitrobot Mark IV (Thermo Fisher Scientific). Cryo-EM imaging was performed on a Titan Krios at 300 kV accelerating voltage in the Center of Cryo-Electron Microscopy, Zhejiang University. Micrographs were recorded using a Gatan K2 Summit direct electron detector in counting mode with a nominal magnification of ×29,000, which corresponds to a pixel size of 1.014 Å. Movies were obtained using serialEM at a dose rate of about 7.8 electrons per Å2 per second with a defocus ranging from −0.5 to −2.5 μm. The total exposure time was 8 s and intermediate frames were recorded in 0.2-s intervals, resulting in an accumulated dose of 62 electrons per Å2 and a total of 40 frames per micrograph. A total of 2,707 and 5,871 movies were collected for the BCM-bound and cortisol-bound GPR97–Go complex, respectively.

Cryo-EM data processing

Dose-fractionated image stacks for the BCM–GPR97–Go complex were subjected to beam-induced motion correction using MotionCor2.144. Contrast transfer function (CTF) parameters for each non-dose-weighted micrograph were determined by Gctf45. Particle selection, 2D and 3D classifications of the BCM–GPR97–Go complex were performed on a binned data set with a pixel size of 2.028 Å using RELION-3.0-beta246.

For the BCM–GPR97–Go complex, semi-automated particle selection yielded 2,026,926 particle projections. The projections were subjected to reference-free 2D classification to discard particles in poorly defined classes, producing 911,519 particle projections for further processing. The map of the 5-HT1BR–miniGo complex (EMDB-4358)47 low-pass filtered to 40 Å was used as a reference model for maximum-likelihood-based 3D classification, resulting in one well-defined subset with 307,700 projections. Further 3D classifications focusing the alignment on the complex produced two good subsets that accounted for 166,116 particles, which were subsequently subjected to 3D refinement, CTF refinement and Bayesian polishing. The final refinement generated a map with an indicated global resolution of 3.1 Å at a Fourier shell correlation of 0.143.

For the cortisol–GPR97–Go complex, particle selection yielded 4,323,518 particle projections for reference-free 2D classification. The well-defined classes with 2,201,933 particle projections were selected for a further two rounds of 3D classification using the map of the BCM-bound complex as reference. One good subset that accounted for 335,552 particle projections was selected for a further two rounds of 3D classifications that focused the alignment on the complex, and produced one high-quality subset with 75,814 particle projections. The final particle projections were subsequently subjected to 3D refinement, CTF refinement and Bayesian polishing, which generates a map with a global resolution of 2.9 Å. Local resolution for both density maps was determined using the Bsoft package with half maps as input maps48.

Model building and refinement

For the structure of the BCM–GPR97–Go complex, the initial template of GPR97 was generated using the module ‘map to model’ in PHENIX44. The coordinate of the 5-HT1BR–Go complex (PDB ID: 6G79) was used to generate the initial models for Go (ref. 44). Models were docked into the EM density map using UCSF Chimera49, followed by iterative manual rebuilding in COOT50 according to side-chain densities. BCM and lipid coordinates and geometry restraints were generated using phenix.elbow. BCM was built to the model using the ‘LigandFit’ module in PHENIX. The placement of BCM shows a correlation coefficient of 0.81, indicating a good ligand fit to the density. The model was further subjected to real-space refinement using Rosetta51 and PHENIX44.

For the structure of the cortisol–GPR97–Go complex, the coordinates of GPR97 and Go from the BCM-bound complex and scFv16 from the human NTSR1–Gi1 complex (PDB ID: 6OS9) were used as initial model. Models were docked into the density map and then were manual rebuilt in COOT. The agonist cortisol was built to the model using the ‘LigandFit’ module as described, showing a good density fit with a correlation coefficient of 0.80. The model was further refined using Rosetta51 and PHENIX44. The final refinement statistics for both structures were validated using the module ‘comprehensive validation (cryo-EM)’ in PHENIX44. The goodness of the fit of the model to the map was performed for both structures using a global model-versus-map FSC (Extended Data Fig. 2). The refinement statistics are provided in Extended Data Table 1. Figures of the structures were generated using UCSF Chimera, UCSF ChimeraX52 and PyMOL53.

Molecular dynamics simulation of the BCM–GPR97 and cortisol–GPR97 complexes

On the basis of the favour binding poses of BCM and cortisol with the receptor GPR97, which was calculated by the LigandFit program of PHENIX, the GPR97–agonist complexes were substrate from the two GPR97–agonist–mGo complexes for molecular dynamics simulation. The orientations of receptors were calculated by the Orientations of Proteins in Membranes (OPM) database. Following this, the whole systems were prepared by the CHARM-GUI and embedded in a bilayer that consisted of 200 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC) lipids by replacement methods. The membrane systems were then solvated into a periodic TIP3P water box supplemented with 0.15 M NaCl. The CHARMM36m Force Filed was used to model protein molecules, CHARMM36 Force Filed for lipids and salt with CHARMM General Force Field (CGenFF) for the agonist molecules BCM and cortisol.

Then, the system was subjected to minimization for 10,000 steps using the conjugated gradient algorithm and then heated and equilibrated at 310.13 K and 1 atm for 200 ps with 10.0 kcal mol−1 Å−2 harmonic restraints in the NAMD 2.13 software. Next followed five cycles of equilibration for 2 ns each at 310.13 K and 1 atm, for which the harmonic restraints were 5.0, 2.5, 1.0, 0.5 and 0.1 kcal mol−1 Å−2 in sequence.

Production simulations were run at 310.13 K and 1 atm in the NPT ensemble using the Langevin thermostat and Nose–Hoover method for 200 ns. Electrostatic interactions were calculated using the particle mesh Ewald (PME) method with a cut-off of 12 Å. Throughout the final stages of equilibration and production, 5.0 kcal mol−1 Å−2 harmonic restraints were placed on the residues of GPR97 that were within 5 Å of Go in the BCM (or cortisol)–GPR97–Go complex to ensure that the receptor remained in the active state in the absence of the G protein. Trajectories were visualized and analysed using Visual Molecular Dynamics (VMD, version 1.9.3)

cAMP ELISA detection in Y-1 cells

Y-1 cells were transfected with Gpr97 siRNA (si-97, GUGCAGGGAAUGUCUUUAA) or control siRNA (si-Con) for 48 h. After starvation for 12 h in serum-free medium, the cells were further stimulated with cortisone (8 nM), forskolin (5 μM) (Sigma-Aldrich) or control vehicle for 10 min. Then, cells were washed three times with pre-cooled PBS and resuspended in pre-cooled 0.1 N HCl containing 500 μM IBMX at a 1:5 ratio (w/v). The samples were neutralized with 1 N NaOH at a 1:10 ratio (v/v) after 10 min. The supernatants were collected after centrifugation of the samples at 600g for 10 min. The supernatants were then prepared for cAMP determination using the cAMP Parameter Assay Kit (R&D Systems) according to the manufacturer’s instruction. The Gpr97 expression level under various conditions were further confirmed using quantitative real-time PCR.

Corticosterone measurements

Mouse adrenocorticotoma cell line Y-1 cells were transfected with Gpr97 siRNA (si-97) or control siRNA (si-Con) for 48 h. Then, the cells were treated with serum-free medium for 12 h. After that, cortisone (16 nM) or ACTH (0.5 μM) were added to cells for 30 min. The supernatants of the cell culture medium were collected for measurements of corticosterone by ELISA according to the manufacturer’s instructions.

Quantitative real-time PCR

Total RNA of cells was extracted using a standard TRIzol RNA isolation method. The reverse transcription and PCR experiments were performed with the Revertra Ace qPCR RT Kit (TOYOBO FSQ-101) using 1.0 μg of each sample, according to the manufacturer’s protocols. The quantitative real-time PCR was conducted in the Light Cycler apparatus (Bio-Rad) using the FastStart Universal SYBR Green Master (Roche). The mRNA level was normalized to GAPDH in the same sample and then compared with the control. The forward and reverse primers for GPR97 used in the experiments were CAGTTTGGGACTGAGGGACC and GCCCACACTTGGTGAAACAC. The mRNA level of GAPDH was used as an internal control. The forward and reverse primers for GAPDH were GCCTTCCGTGTTCCTACC and GCCTGCTTCACCACCTTC.

cAMP inhibition assay

To measure the inhibitory effects on forskolin-induced cAMP accumulation of different GPR97 constructs or mutants in response to different ligands or constitutive activity, the GloSensor cAMP assay (Promega) was performed according to previous publications12,13. HEK293 cells were transiently co-transfected with the GloSensor and various versions of GPR97 or vehicle (pcDNA3.1) plasmids using PEI in six-well plates. After incubation at 37 °C for 24 h, transfected cells were seeded into 96-well plates with serum-free DMEM medium (Gibco) and incubated for another 24 h at 37 °C in a 5% CO2 atmosphere. Different ligands were dissolved in DMSO (Sigma) to a stock concentration of 10 mM and followed by serial dilution using PBS solution immediately before the ligand stimulation. The transfected cells were pre-incubated with 50 μl of serum-free DMEM medium containing GloSensor cAMP reagent (Promega). After incubation at 37 °C for 2 h, varying concentrations of ligands were added into each well and followed by the addition of forskolin to 1 μM. The luminescence intensity was examined on an EnVision multi-label microplate detector (Perkin Elmer).

The Gqo protein activation BRET assay

According to previous publications, the BCM dipropionate-induced GPR97 activity could be measured by chimeric Gqo protein assays25. The Gqo BRET probes were generated by replacing the six amino acids of the C-terminal of Gq-RlucII with those from GoA1, creating a chimeric Gqo-RlucII subunit47. GFP10 was connected to Gγ. The Gqo protein activation BRET assay was performed as previously described54. In brief, HEK293 cells were transiently co-transfected with control D2R and various GPR97 constructs, plasmids encoding the Gqo BRET probes, incubated at 37 °C in a 5% CO2 atmosphere for 48 h. Cells were washed twice with PBS, collected and resuspended in buffer containing 25 mM HEPES, pH 7.4, 140 mM NaCl, 2.7 mM KCl, 1 mM CaCl2, 12 mM NaHCO3, 5.6 mM d-glucose, 0.5 mM MgCl2 and 0.37 mM NaH2PO4. Cells that were dispensed into a 96-well microplate at a density of 5–8 × 104 cells per well were stimulated with different concentrations of ligands. BRET2 between RLucII and GFP10 was measured after the addition of the substrate coelenterazine 400a (5 μM, Interchim) (Cayman) using a Mithras LB940 multimode reader (Berthold Technologies). The BRET2 signal was calculated as the ratio of emission of GFP10 (510 nm) to RLucII (400 nm).

Measurement of receptor cell-surface expression by ELISA

To evaluate the expression level of wild-type GPR97 and its mutants, HEK293 cells were transiently transfected with wild-type and mutant GPR97 or vehicle (pcDNA3.1) using PEI regent at in six-well plates. After incubation at 37 °C for 18 h, transfected cells were plated into 24-well plates at a density of 105 cells per well and further incubated at 37 °C in a 5% CO2 atmosphere for 18 h. Cells were then fixed in 4% (w/v) paraformaldehyde and blocked with 5% (w/v) BSA at room temperature. Each well was incubated with 200 μl of monoclonal anti-FLAG (F1804, Sigma-Aldrich) primary antibody overnight at 4 °C and followed by incubation of a secondary goat anti-mouse antibody (A-21235, Thermo Fisher) conjugated to horseradish peroxide for 1 h at room temperature. After washing, 200 μl of 3,3′,5,5′-tetramethylbenzidine (TMB) solution was added. Reactions were quenched by adding an equal volume of 0.25 M HCl solution and the optical density at 450 nm was measured using the TECAN (Infinite M200 Pro NanoQuant) luminescence counter. For determination of the constitutive activities of different GPR97 constructs or mutants, varying concentrations of desired plasmids were transiently transfected into HEK293 cells and the absorbance at 450 nm was measured.

The FlAsH-BRET assay

HEK293 cells were seeded in six-well plates after transfection with GPR97-FlAsH with Nluc inserted in a specific N-terminal site. Before the BRET assay, HEK293 cells were starved with serum for 1 h. Then cells were digested, centrifuged and resuspended in 500 μl BRET buffer (25 mM HEPES, 1 mM CaCl2, 140 mM NaCl, 2.7 mM KCl, 0.9 mM MgCl2, 0.37 mM NaH2PO4, 5.5 mM d-glucose and 12 mM NaHCO3). The FlAsH-EDT2 was added at a final concentration of 2.5 μM and incubated at 37 °C for 60 min. Subsequently, HEK293 cells were washed with BRET buffer and then distributed into black-wall clear-bottom 96-well plates, with approximately 100,000 cells per well. The cells were treated with a final concentration of BCM and cortisol at 10−5 to 10−11 and then coelenterazinc H was added at a final concentration of 5 μM, followed by checking the luciferase (440–480 nm) and FlAsH (525–585 nm) emissions immediately. The BRET ratio (emission enhanced yellow fluorescent protein/emission Nluc) was calculated using a Berthold Technologies Tristar 3 LB 941 spectrofluorimeter. The procedure was modified from those described previously34,55,56.

Statistical analysis

A one-way ANOVA test was performed to evaluate the statistical significance between various versions of GPR97 and their mutant in terms of expression level, potency or efficacy using GraphPad Prism. For all experiments, the standard error of the mean of the values calculated based on the data sets from three independent experiments is shown in respective figure legends.

Reporting summary

Further information on research design is available in the Nature Research Reporting Summary linked to this paper.


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Bio Chemistry

A ‘Build and Retrieve’ methodology to simultaneously solve cryo-EM structures of membrane proteins

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    Bio Chemistry

    Molecular basis for RNA polymerase-dependent transcription complex recycling by the helicase-like motor protein HelD

    Republished by Plato



    Structure determination of the transcription elongation complex

    Bacillus subtilis is the model representative organism of the medically and industrially important Firmicutes phylum that have genomes with a low G + C content. Despite considerable effort, no structure of RNAP from the low G + C Gram positives has been determined to date. RNAP core (α2ββω) was purified (Supplementary Fig. 1), and it’s activity established using a HelD-dependent stimulation of multi-round transcription assay which gave identical results to those observed in previous studies12 (Fig. 1a). We then used cryo-EM to determine the structure of the B. subtilis RNAP transcription elongation complex (EC) at 3.36 Å, to enable an understanding of the conformational changes caused by HelD during transcription complex recycling (Fig. 1b, c, Table 1; Supplementary Fig. 2, Movie 1). RNAP in the Firmicutes is the smallest multi-subunit polymerase17,18, and given the industrial and clinical importance of this group of bacteria this complex will serve as an invaluable reference structure.

    Fig. 1: Structure of RNAP elongation complex.

    a Left-hand graph shows multi-round transcription assays of purified RNAP core supplemented with increasing amounts of HelD (as a ratio with RNAP), HelD only (0:32, negative control) and the native RNAP-HelD complex purified from B. subtilis (RNAP-HelD). Black dots represent data obtained using wild-type HelD, magenta dots, data obtained using HelD K239A. The right-hand graph shows ATPase assays of wild-type HelD, HelD K239A, and BSA (negative control). Error bars, SD. Each experiment was performed three times in technical duplicate. Source data are provided in the Source Data files. b, c Cryo-EM reconstruction of the RNAP elongation complex (EC). The electron density map is semi-transparent in the same colours as the subunits in the cartoon structure: αI beige, αII brown, β azure, β’ yellow, ω light green, template DNA purple, non-template DNA pink, and RNA orange. The dotted circles show the location of the βln5 insertion, ε binding site (b) and β E696-G705 insertion (c), respectively. The upstream and downstream sides of RNAP are indicated in c. d Top shows a schematic of the nucleic acids coloured as in panels b and c. +1 represents the template DNA nucleotide positioned within the active site. + integers represent nucleotides base paired as DNA on the downstream side of RNAP, and −ve integers represent the DNA-RNA hybrid on the upstream side. The arrow indicates the position of the unpaired +1 nucleotide in the active site of RNAP in the lower part of the panel. Intermolecular bonds (polar and non-polar) are indicated by the dashed lines. Relevant template (t) and non-template (nt) nucleotides are numbered appropriately and amino acids shown with the RNAP subunit as a prefix. The catalytic Mg2+ ion is shown as a green sphere to indicate the active site. e An enlarged view of the upstream side of the EC close to the entry of the RNA exit channel. The salt bridge formed between β R800 and β’ D245 is shown as black dashed line. The space below the β’ lid that could accommodate a 9th DNA-RNA base pair is shown as a dotted rectangle, with the β’ rudder helping guide RNA towards the β’ lid and the RNA exit channel.

    Table 1 Cryo-EM data collection, refinement and validation statistics.

    Despite nuclease treatment of the cell lysate, upon 3D reconstruction of the core structure, nucleic acid was clearly visible indicating that throughout the purification process the core enzyme remained tightly bound to nucleic acids which protected them from nuclease treatment (Supplementary Table 1). Therefore, the structure presented represents an elongation complex (EC) with non-specified nucleic acid sequence (i.e., the reconstructed density shows well-defined ribose-phosphate groups with an average of random base sequences). Typically, the subunit composition of core bacterial RNAP is represented as α2ββʹω, but in previous work we identified an additional small subunit called ε was present in B. subtilis RNAP in addition to ω19. However, holoenzyme preparations from the strain (LK637, Δ δ; Methods) used in this study lacked ε (Supplementary Fig. 1a) and so the core structure is presented lacking this subunit (although ε is present in the structure of RNAP core in complex with HelD, see below). Previous studies have also shown deletion of ε causes no detectable phenotype or change in gene expression profiles, and RNAP core preparations lacking ε have indistinguishable activity compared to those that do contain it19,20. Comparison of EC and RNAP-HelD complexes showed no significant structural differences in the region where ε binds and so in Fig. 1b the ε binding site is indicated as a dotted circle, and in Supplementary Fig. 4a ε is shown as it is clear that RNAP isolated from B. subtilis is a heterogeneous mixture of core (α2ββ’ω) ± δ, ε, and HelD, in addition to multiple different σ factors21.

    The EC is 150 Å × 112 Å × 123 Å (L × W × H, Table 1), and is broadly comparable to the dimensions of core/elongation complexes from other species (157 × 153 × 136 Å; E. coli 6ALF, 183 × 107 × 115 Å; Mycobacterium smegmatis 6F6W, and 170.1 × 110.1 × 127.8 Å; Thermus thermophilus 2O5I)22,23,24, although it appears to be more slender and elongated than the roughly globular E. coli, and shorter than the M. smegmatis and T. thermophilus enzymes due to the lack of insertion sequences (Supplementary Fig. 4).

    Due to the high level of sequence conservation amongst RNAPs, the overall structure of the EC was similar to those from other organisms and largely consistent with homology models used in previous work on structure/function studies with B. subtilis RNAP25,26,27,28. However, modelling had been unable to establish the structure of the ~180 amino acid βln5 insertion in the β2 lobe. The β2 lobe is one of the least well-conserved regions of bacterial RNAPs, and is a hot-spot for the presence of lineage-specific insertions17 (Supplementary Fig. 4a). The only other region that was significantly different to other bacterial RNAPs was a 10 amino acid loop from β E696-G705 that protrudes from the bottom of the enzyme (Fig. 1c, Supplementary Fig. 4a). Refinement and building sequence into the resulting density indicated the B. subtilis β2 lobe is a continuous globular structure and that the βln5 insertion increases the size asymmetry between the β lobes compared to other Gram positive RNAPs such as those from M. smegmatis and M. tuberculosis23,29 (Fig. 1b, Supplementary Fig. 4). Searches using DALI30 found no structural matches to the βln5 insertion leaving its function similarly enigmatic to those of most other lineage-specific insertions.

    The absence of lineage-specific insertions perhaps helps to account for the additional subunits found in the Firmicutes such as δ and ε, and this and the accompanying paper by Pei et al.31 identify the location of these subunits. This suggestion is potentially supported by examination of the T. thermophilus structure around its βln10 and βln12 insertions that localise to a region very close to the ε binding site. Superimposition of ε into the T. thermophilus EC structure shows steric clashes between ε and the insertions (Supplementary Fig. 4c) raising the possibility that they serve similar functions. The location of ε also corresponds to that of a domain of archaeal and eukaryotic Rpo3/RPB3 subunits associated with enzyme stability (boxed insert, Supplementary Fig. 4a) and it is interesting to note that both B. subtilis (able to grow up to ~52 °C) and the thermophile T. thermophilus (up to ~79 °C) have structural elements/subunits located in this area that links the α2, β, and β’ subunits whereas the mesophilic E. coli and M. smegmatis do not.

    The ω subunit in B. subtilis is 67 amino acids vs the 80 amino acid length of E. coli ω. The main structural difference appears to be in the lack of a C-terminal α-helix which is prominent in E. coli RNAP, but lacking in B. subtilis and Mycobacterial structures. As with all other RNAP core and EC structures solved to date, the C-terminal domains of the α subunits were not visible due to the flexible linker connecting the N- and C-terminal domains.

    Detailed examination of the elongation complex also revealed important features associated with mechanistic aspects of RNA synthesis. The density for fork-loop 2 (FL2) is well defined, consistent with its role in DNA strand separation on the downstream edge of the transcription bubble. The EC active site is similar in structure to that reported previously for the T. thermophilus and E. coli ECs22,24 (2O5I, and 6ALF, respectively) and is in a post-translocation conformation with the 3′ end of the RNA transcript adjacent to the +1 site, with an unbent bridge-helix (BH) and the trigger-loop (TL) in the open conformation (Fig. 1d). This conformation is consistent with an elongation complex primed to receive an incoming NTP via the secondary channel.

    FL2 residue β R498 interacts with the ribose and phosphate moieties of the final base in the non-template DNA strand prior to strand separation and formation of the transcription bubble and likely acts to facilitate formation of the downstream edge of the transcription bubble (Fig. 1d). The template base in the +1 site is held in position for base-pairing with the incoming substrate NTP through interaction with the highly conserved T794 and A795 of the BH, and may also be stabilised through stacking with the base in the −1 position (Fig. 1d). β R496 of FL2 interacts with the phosphodiester backbone of RNA bases 4 and 5 of the new transcript (Fig. 1d). In addition, residues Q469, P520, E521, N524, I528, K924 and K932 of the rifampicin binding pocket of the β subunit form numerous interactions with the newly formed transcript (RNA residues 1–5) as has been previously reported32,33. The salt bridge between β R800 and β’ D245 that closes the primary channel off from the RNA exit channel34,35 is clearly visible confirming that the elements on the upstream side of the transcription bubble, the rudder and lid, that are responsible for facilitating reannealing of the template and non-template strands and guiding RNA into the exit channel are in positions consistent with these assigned roles (Fig. 1e).

    Electron density for RNA beyond the 8th nucleotide is poor, preventing further mapping of the transcript up to and through the exit channel. Likewise, density for DNA on the upstream side is poorly defined consistent with conformational flexibility in this region of RNAP26. Structural modelling, and comparison with ECs from other organisms22,24, is consistent with there being sufficient space for a transcription bubble comprising a 9 bp template DNA-RNA hybrid prior to upstream DNA strand re-annealment and entry of the transcript into the exit channel guided by hydrophobic interaction with conserved β’ lid residues V242 and L244 (dotted box, Fig. 1e). The 9th RNA-DNA base pair has likely been degraded by nuclease activity during preparation of the complex. Overall, this structure serves as a valuable resource for structure-function studies with RNAP from the Firmicutes as well as being a reference structure to enable full understanding of the conformational changes involved in transcription complex recycling induced upon binding to HelD (below).

    The structure of an RNAP-HelD transcription recycling complex

    RNAP-HelD complexes were isolated from a culture of B. subtilis carrying a deletion of the rpoE gene that encodes the δ subunit, shown previously to act synergistically with HelD12 (Supplementary Fig. 1). HelD itself is required for transcription complex recycling, and can perform this function independently of δ12 which is absent in many organisms that contain genes encoding HelD proteins (e.g. Clostridia). The purified complex stimulated transcription ~2-fold, similar to that observed with in vitro assembled complexes12, establishing its biological activity (Fig. 1a).

    We determined the structure of the RNAP–HelD complex using single particle cryo-electron microscopy (cryo-EM) to 3.36 Å resolution (Supplementary Fig. 3), followed by atomic modelling (Fig. 2a, Table 1; Supplementary Movie 2). The resulting structure revealed that HelD, which is located on the downstream side of RNAP, has two arm domains that penetrate deep into the primary and secondary channels of RNAP (clamp arm; CA, and secondary channel arm; SCA, respectively, Fig. 2a–c), which account for the strong HelD-RNAP interaction12,13,36. The native RNAP-HelD preparation also contained the RNAP ε subunit19 and showed it bound on the downstream side of RNAP in a concave space between the two α, β, and β’ subunits (Fig. 2a; see Supplementary Fig. 4).

    Fig. 2: Structure of RNAP in complex with HelD.

    ac Different views of the cryo-EM reconstruction of the RNAP-HelD complex. The electron density map is semi-transparent in the same colours as in Fig. 1 with the addition of ε green, and HelD red. The HelD clamp arm (CA), secondary channel arm (SCA), 1 A Torso, and 2 A Head domains are labelled, as are the up- and downstream sides of RNAP (a). The primary channel that is formed between the β and β’ subunits is shown in b, and the secondary channel indicated by the dotted circle in a. The change in orientation between the views in panels a and b is indicated by the arrows, with the right side arrow indicating a rotation of the view in a upwards from the α2/ε end that would move the β’ end down. The left arrow indicates the subsequent 165° clockwise rotation of the complex to give the view in b. The view in c is a simple 90° rotation of the orientation in b.

    HelD itself has an unusual 4-domain structure (Fig. 3a, b). The first 203 amino acids (aa) form the secondary channel arm (SCA), which is joined to a super-family 1 (SF1) 1 A domain (aa 204–291 and 539–610). In SF1 helicases, domain 1 A is split by the insertion of a 1B domain associated with helicase function37, but in HelD it is split by the clamp arm (CA; aa 292–538). Residues 610–774 form a continuous SF1 2 A domain, which is usually split by a 2B insertion in SF1 helicases, that represents the ‘head’ of HelD. The overall appearance of the protein is that of a torso and head (domains 1 A and 2 A, respectively) flanked by a pair of muscular arms (SCA and CA), giving it a rather thuggish appearance (Fig. 3b, c).

    Fig. 3: Structure and sequence conservation of HelD.

    a Linear representation of the structure of HelD with a map of coloured domains and conserved sequence motifs (top), and a histogram of sequence identity (bottom). b The structure of HelD (bottom) coloured according to the schematic in panel a. SCA is shown in yellow, SF1 helicase-like domain 1 A in red, CA in dark grey, and SF1 helicase-like domain 2 A in orange. Conserved sequence motifs are shown in purple, with the Walker A/B ATP-binding site in cyan. c, shows a surface-rendered impression of HelD with the same colouring as in b. The bottom panel shows an expanded view of the boxed region in the top panel with a semi-transparent surface and conserved amino acids that form the ‘Trp cage’ shown as purple sticks, and the conserved Trp in dark green.

    Although HelD is widely distributed amongst Gram-positive bacteria, the distinctive arm domains represent the regions of lowest sequence conservation despite being responsible for the majority of interactions with RNAP as well as for its transcription recycling activity13 (Fig. 3a, Supplementary Figs. 57). It is also clear that there are at least two distinct classes of HelD (Classes I and II, Supplementary Figs. 5, 6, Supplementary Table 2); Class I is represented by the B. subtilis protein and is present in the low G + C Gram positives, whereas Class II is represented by the M. smegmatis protein (see accompanying paper by Kouba et al.38), and is present in the high G + C Gram-positives. Some organisms contain multiple copies of HelD (e.g. Lactobacillus plantarum, Class I; Nonomuraea wenchangensis, Class II; Supplementary Fig. 5), and even within the same organism, sequence conservation between the copies is relatively low in the SCA and CA domains (Supplementary Fig. 8). Previous studies showed that HelD in which the SCA (aa 1–203) had been deleted was still capable of binding RNAP, hydrolysing ATP, and binding DNA, but not transcription recycling13. These observations suggest that the function of the arm domains is centred around mechanical work rather than the formation of highly-conserved functionally-significant interprotein interactions.

    Despite the clear separation into two classes, sequence alignment allowed the identification of conserved motifs common to all HelD proteins (Fig. 3a; Supplementary Table 2). The transcription recycling function of HelD is dependent on ATP hydrolysis12, with ATP-binding motifs located in the 1 A (torso) domain (cyan residues, Fig. 3a, b). Alteration of the absolutely conserved K239 to A in the Walker A motif resulted in the complete loss of transcription recycling and ATPase activity (Fig. 1a). The remaining conserved motifs form a network of interactions that are mainly centred in the region between the SCA and 1 A domains, with the absolutely conserved residue W137 in a hydrophobic pocket between them (purple residues, Fig. 3a, c). These extensive interactions anchor the SCA to the 1 A domain, helping to couple ATP hydrolysis to mechanical movement of the CA (see below).

    HelD causes major conformational changes in RNAP

    Comparison of the core elements of the EC and RNAP-HelD structures (α2ββ’ω subunits) shows HelD causes a major conformational change mainly due to the opening of the β’ clamp by the CA, with very little change elsewhere (Fig. 4a, b; Supplementary Movie 3, and see below). PISA39 was used to analyse protein-protein contacts in the RNAP-HelD and elongation complexes (Supplementary Table 3). Complexation with HelD reduces the contact area between RNAP subunits β and β’ by over 6% while other contact areas remain similar, consistent with the extensive conformational change caused to the EC upon binding of HelD.

    Fig. 4: HelD-induced conformational change in RNAP.

    a, b Cryo-EM electron density maps (α2ββω subunits only) of RNAP EC (a) and RNAP–HelD complex (b). HelD density has been removed for clarity, and nucleic acids from the E. coli EC (PDB ID 6ALF) superimposed over the B. subtilis EC nucleic acids as a visual aid for the scale of conformational changes that occur in the primary DNA-binding and RNA exit channels upon binding HelD. Nucleic acids are coloured the same as in Fig. 1. The longer RNA from the E. coli EC helps illustrate the increase in aperture of the RNA exit channel. c left side EC, right side RNAP-HelD complex with HelD removed for clarity, with β (cyan) and βʹ (purple) residues shown to illustrate the change in aperture of the RNA exit channel β R800 and βʹD245 and DNA binding channel β P242 and βʹN283. Black arrow in left panel indicates the region of the βʹ clamp that is contacted by the HelD clamp arm and the double-ended arrow in the right panel indicates the movement of subunits away from each other on HelD binding.

    As part of transcription complex recycling, the elongating RNA as well as the DNA template needs to dissociate from RNAP. RNA passes through the exit channel on the upstream side of RNAP. There was no major conformational change to elements at the entry of the exit channel other than those that are translocated as part of the opening of the β’ clamp (Fig. 4). The translocation of the β’ clamp results in breaking of the conserved salt bridge between β R800 and β’ D245 that is important in guiding RNA into the exit channel34,35, increasing the width of the aperture from 11 to 20 Å (αc–αc; Fig. 4c, Supplementary Movie 3). This separation, along with widening of the primary channel, facilitates RNA exit from the complex.

    The most dramatic effect of HelD on RNAP is the widening of the primary channel from 21 to 47 Å between β2 lobe P242 and β’ clamp helix N283, that would cause a loss of contact with DNA in the primary channel, enabling recycling of RNAP (Fig. 4b, c). This is facilitated by the proximity of the CA to the SW5 region of the β’ clamp, that acts as a hinge during clamp movement18,40 (Supplementary Fig. 9, Movie 3).

    Detailed examination of the SCA and CA interactions with RNAP enabled us to define the molecular events that occur during transcription complex recycling. Images of the active site region in the EC (Fig. 5a), RNAP-HelD complex (Fig. 5b) and an overlay of the two views (Fig. 5c) shows how HelD SCA insertion via the secondary channel causes distortion of the bridge-helix and trigger-loop as well as steric clashes with nucleic acids. The prokaryotic Gre factors, DksA, and eukaryotic TFIIS are known to bind in the secondary channel of RNAP via a pair of anti-parallel α helices/hairpin loop41,42,43. The acidic tips of these proteins reside close to, but on the downstream side of the catalytic Mg2+. The SCA of HelD bears superficial similarity to GreB/DskA, but is longer and the tip extends past the catalytic Mg2+ (Supplementary Fig. 10). The acidic tip (D56 and D57) will electrostatically repel the transcript upon penetration of the SCA into the active site, with the SCA causing significant steric clashes with the transcript and template DNA strand when fully inserted (Fig. 5c). The bridge-helix and trigger-loop, are dynamic structures that play a key role in the transcription cycle44; the entry of the SCA into the secondary channel causes partial folding of the open trigger-loop conformation observed in the EC structure and a major distortion of the bridge-helix that would sterically clash with the template DNA in the active site (Fig. 5a–c; Supplementary Movie 3). Thus, the SCA tip itself, in combination with the distortion its insertion causes in the bridge-helix, will result in physical displacement of template DNA and RNA from the active site of RNAP, facilitated by electrostatic repulsion between the acidic SCA tip residues and the transcript.

    Fig. 5: HelD interactions with RNAP.

    a View of the active site region of the EC with the bridge-helix in teal, trigger-loop in yellow, template DNA strand in purple and RNA in orange. b The same view of the active site region of the RNAP-HelD complex with the SCA of HelD shown in red with the acidic D56 and D57 residues shown as sticks. c An overlay of the regions shown in a and b with the insertion of the SCA into the secondary channel indicated by the red arrow. EC elements are shown semi-transparent with nucleic acids coloured according to the scheme in Fig. 1. The bridge-helix and trigger-loop elements that are distorted by the SCA are shown, with the direction of movement from EC to HelD complex indicated by the black arrow. The catalytic Mg2+ ion is shown as a green sphere. d Hydrogen-bond and salt-bridge interactions (dashed blue lines) between the tip of the SCA (red semi-transparent cartoon with purple sticks) and RNAP. Subunit colouring is the same as in Fig. 1. The conserved active site Asp residues that chelate the catalytic Mg2+ (green sphere, grey dashed lines) are also shown to illustrate how the tip of the SCA cages but does not interact directly with residues involved in RNAP catalysis. The enlarged box on the right corresponds to the boxed region shown for the whole complex on the left.

    The fully inserted tip of the SCA is in close proximity to the absolutely conserved active-site β’447NADFDGD453, forming a network of interactions around this motif, but does not directly interact with either the catalytic Mg2+ or the Asp residues that coordinate it (Fig. 5d, Supplementary Table 4). Thus, upon dissociation of HelD, the core RNAP would be competent for re-use in transcription, as seen in the transcription recycling assays in Fig. 1a. Finally, insertion of the SCA into the secondary channel would block NTP entry into the active site.

    The salt-bridge and H-bond contacts the CA makes with the β’ clamp are listed in Supplementary Table 4, but the bulk of interactions are made by hydrophobic residues with little sequence conservation between even closely-related genera (Supplementary Table 5, Supplementary Fig. 6a). This region is the location of an insertion that spans across the primary channel towards the active site in Class II HelD proteins (see accompanying paper by Kouba et al.38; Supplementary Fig. 6b). The tip of this insertion has a similar location to the tip of the SCA of B. subtilis Class I HelD and is also acidic, suggesting electrostatic repulsion of nucleic acids is also important in the activity of Class II HelDs. In our Class I HelD structure, the site of this insertion is close to an area of density in the cryo-EM reconstruction that at low threshold values could be consistent with the presence of nucleic acid (Supplementary Fig. 11). Examination of the surface charge of HelD revealed a region of high overall positive charge on the inside of the CA. Superposition with the nucleic acids from the EC show that this positively-charged patch is in a position where it could interact with the downstream dsDNA (Supplementary Fig. 11a), consistent with nucleic acid-binding data13. It is also possible that this patch may be important for interaction with the unstructured negatively-charged C-terminal domain of δ which acts synergistically with HelD during transcription complex recycling12 (see accompanying paper by Pei et al.31). The end of the CA forms a relatively flat ~320 Å2 surface that acts as a platform to push up against the β’ clamp, resulting in loss of contact with the DNA bound in the EC (Figs. 2a–c, 4, Supplementary Movie 3). Therefore, the purpose of the CA appears to involve the opening of the β’ clamp through brute force rather than by the formation of a specific network of conserved interactions.

    Movement of the clamp arm of HelD drives conformational change in RNAP

    Closer examination of the RNAP–HelD complex using 3D variability analysis (3DVA)45 allowed identification of regions of conformational flexibility that underpin the dynamic processes of HelD activity in transcription recycling. Overall, the region behind SW5 towards the α dimer, including the secondary channel and SCA of HelD, showed little or no

    conformational variability, but the primary channel encompassing elements of the β1 and 2 lobes and the β’ clamp did (Fig. 6a, Supplementary Movie 4).

    Fig. 6: 3D variability analysis of the RNAP–HelD complex and a model for HelD-catalysed RNAP recycling.

    a, b Different orientations of the most distinct conformations determined by 3DVA. HelD is shown in solid colours with RNAP semi-transparent. The red conformation of HelD is matched to the pale green conformation of RNAP, and the purple HelD with the yellow RNAP. Orange arrows indicate the movement of the juxtaposed region of HelD, and cyan arrows regions of RNAP. Structural elements of RNAP referred to in the text are labelled as well as the up- (UP) and downstream (DOWN) sides of RNAP. c Model for HelD-catalysed recycling. HelD is shown in red, RNAP in white, DNA in purple (template) and pink (non-template), and RNA in orange. Clockwise from top left; HelD locates a stalled EC and binds with the SCA penetrating the secondary channel and the CA moving into position on the β’ clamp. The SCA is wedged deep within RNAP, and through conserved interactions with the 1 A torso domain, locks HelD in position. The contacts the CA makes with the β’ clamp, open the DNA-binding and RNA-exit channels to enable dissociation of nucleic acids from the EC, possibly assisted by interaction of the DNA with the positively-charged patch on the CA of HelD. HelD (and nucleic acid) dissociation is facilitated by the conformational changes facilitated by ATP binding/hydrolysis (ATP flash). Finally, core RNAP that has been released from the complex is free to enter a fresh round of transcription.

    The 3D variability analysis indicates that HelD causes the β’ clamp to open and twist so that the downstream side of RNAP opens slightly (curved cyan arrow, Fig. 6a). At the same time, the β2 lobe moves up (straight cyan arrow, Fig. 6a) along with a slight twisting of the β1 lobe and β flap (Fig. 6a, Supplementary Movie 4). With respect to HelD, there was no change in the SCA tip adjacent to the RNAP active site, but there was lateral movement of the portion located outside the secondary channel towards the β’ jaw (Fig. 6b). This resulted in little, if any, conformational change in the hydrophobic ‘cage’ surrounding the conserved W137 residue. Accordingly, there was relatively little change in the torso (1 A) domain and ATP-binding site, but the head (2 A) domain moved away from the downstream side of RNAP (curved and straight orange arrows Fig. 6a, b, respectively). The CA of HelD rises up and out slightly, causing the upward twist on the downstream side of the β’ clamp (orange arrow, Fig. 6a; Supplementary Movie 4). Therefore, the results of the 3D variability analysis are consistent with the SCA acting as a wedge that permits conformational change through movement of the CA. The CA is located in a position equivalent to an SF1 helicase 1B domain that utilises ATP hydrolysis to undergo conformational changes required for helicase/translocase activity46,47 and ATP binding/hydrolysis is required for release of HelD (see accompanying papers by Kouba et al. and Pei et al.31,38), most likely due to movement of the CA arm, consistent with the observed conformational flexibility in this region.

    In our structure, and those of the accompanying papers by Pei et al. and Kouba et al., no density for any NTP could be detected in the ATP binding site, even on addition of ATP or non-hydrolysable analogues. In order to bind ATP, domains 1 A (torso) and 2 A (head) need to rotationally open as observed for SF1 helicases48. Our 3DVA suggests this is most likely via movement of the 2 A (head) domain (Fig. 6a, b). However, the sequence from F183-G190 linking the SCA to the 1 A (torso) domain sterically blocks access to the ATP binding site and this ‘gate’ region will also need to open to allow ATP binding and subsequent ADP release (Supplementary Fig. 12). There is no intramolecular bonding between residues T185-I189 and either the SCA or IA (torso) domain, and this may provide the necessary flexibility for gate opening and closing. Given that the ATP binding site was not accessible in all of the structures that are forcing open the DNA binding clamp of RNAP, gate opening may be an event that occurs on conformational change of the CA during nucleic acid release and RNAP recycling.


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