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Melanoma specimens were obtained with informed consent from all patients according to protocols approved by the Institutional Review Boards of the University of Michigan Medical School (IRBMED approvals HUM00050754 and HUM00050085; see ref. 27) and the University of Texas Southwestern Medical Center. Tumours were dissociated in Sterile Closed System Tissue Grinders (SKS Science) in enzymatic digestion medium containing 200 U ml−1 collagenase IV (Worthington) for 20 min at 37 °C. DNase (50–100 U ml−1) was added to reduce clumping of cells during digestion. Cells were filtered with a 40- μm cell strainer to obtain a single-cell suspensions. All melanomas used in this study stably expressed DsRed and luciferase so that the melanoma cells could be unambiguously distinguished from mouse cells by flow cytometry and by bioluminescence imaging. When isolated by flow cytometry, cells were also stained with antibodies against mouse CD45 (30-F11-APC, eBiosciences), mouse CD31 (390-APC, Biolegend), Ter119 (TER-119-APC, eBiosciences) and human HLA-A, -B, -C (G46-2.6-FITC, BD Biosciences) to select live human melanoma cells and to exclude contaminating mouse endothelial and haematopoietic cells. Live human melanoma cells were thus isolated by flow cytometry by sorting cells that were positive for DsRed and HLA and negative for mouse CD45, Ter119 and CD31. All antibody labelling was performed for 20 min on ice, followed by washing and centrifugation. Before flow cytometric analysis, cells were re-suspended in staining medium (L15 medium containing bovine serum albumin (1 mg ml−1), 1% penicillin/streptomycin, and 10 mM HEPES, pH 7.4) containing 4′,6-diamidino-2-phenylindole (DAPI; 5 μg ml−1; Sigma) to eliminate dead cells from sorts and analyses. Sorts and analyses were performed using a FACSAria flow cytometer (Becton Dickinson). After sorting, an aliquot of sorted melanoma cells was always reanalysed to check for purity, which was usually greater than 95%. For analysis of circulating melanoma cells, blood was collected from each mouse by cardiac puncture with a syringe pretreated with citrate-dextrose solution (Sigma). Red blood cells were precipitated by Ficoll sedimentation according to the manufacturer’s instructions (Ficoll Paque Plus, GE Healthcare). Remaining cells were washed with Hanks’ balanced salt solution (Invitrogen) before antibody staining and flow cytometric analysis. For limiting dilution analysis, cells for each mouse were sorted into individual wells of 96-well V-bottomed plates containing staining medium and loaded into syringes directly from the well (one well into one syringe into one mouse). After sorting, cells were counted and resuspended in staining medium with 25% high-protein Matrigel (product 354248; BD Biosciences). Subcutaneous injections were performed into the right flank of NOD.CB17-Prkdcscid Il2rgtm1Wjl/SzJ (NSG) mice (Jackson Laboratory) in a final volume of 50 μl. Each mouse was transplanted with 100 melanoma cells unless otherwise specified. Tumour formation was evaluated regularly by palpation of the injection site, and the subcutaneous tumours were measured every 10 days until any tumour in the mouse cohort reached 2.5 cm in its largest diameter. Mice were monitored daily for signs of distress and euthanized when they exhibited distress according to a standard body condition score or within 24 h of their tumours reaching 2.5 cm in largest diameter, whichever came first. We adhered to this limit in all experiments. Organs were analysed visually and by bioluminescence imaging (see details below) for presence of macrometastases and micrometastases. These experiments were performed according to protocols approved by the Institutional Animal Care and Use Committee at the University of Texas Southwestern Medical Center (protocol 2011-0118). Intravenous injections were done by injecting cells into the tail vein of NSG mice in 100 μl of staining medium. For intrasplenic injections the mice were anaesthetized with isoflourane, then the left flank was shaved and disinfected with an ethanol wipe and iodine swab. An incision was made into the intraperitoneal cavity. The spleen was exposed with forceps and cells were injected slowly in a 40 μl volume of staining medium. The peritoneum was then sutured and skin was closed with clips. Mice were injected with buprenex before surgery and then again 12 h after surgery. A bicistronic lentiviral construct carrying dsRed2 and luciferase (dsRed2-P2A-Luc) was generated (for bioluminescence imaging) and cloned into the FUW lentivrial expression construct. The primers that were used for generating this construct were: dsRed2 forward, 5′-CGACTCTAGAGGATCCatggatagcactgagaacgtc-3′ (capital letters indicate homology to FUW backbone); dsRed2 reverse, 5′-TCCACGTCTCCAGC CTGCTTCAGCAGGCTGAAGTTAGTAGCTCCGCTTCCctggaacaggtggtggc-3′ (capital letters indicate P2A sequences); luciferase forward, 5′-GCCTGCTGAAGCAGGCTGGAGACGTGGAGGAGAACCCTGGACCTGGATCCatggaagacgccaaaaacataaag-3′ (capital letters indicate P2A sequences) and luciferase reverse, 5′-GCTTGATATCGAATTCttacacggcgatctttccgc-3′ (capital letters indicate homology to FUW backbone). All constructs were generated using the In-Fusion HD cloning system (Clontech) and sequence verified. For virus production, 0.9 μg of the appropriate plasmid and 1 μg of helper plasmids (0.4 μg pMD2G and 0.6 μg of psPAX2) were transfected into 293T cells using polyjet (Signagen) according to the manufacturer’s instructions. Replication incompetent viral supernatants were collected 48 h after transfection and filtered through a 0.45- μm filter. Approximately 300,000 freshly dissociated melanoma cells were infected with viral supernatants supplemented with 10 μg ml−1 poybrene (Sigma) for 4 h. Cells were then washed twice with staining medium, and about 25,000 cells (a mixture of infected and non-infected cells) were suspended in staining medium with 25% high-protein Matrigel (product 354248; BD Biosciences) then injected subcutaneously into NSG mice. After growing to 1–2 cm in diameter, tumours were excised and dissociated into single-cell suspensions, and luciferase-dsRed+ or green fluorescent protein (GFP)+ cells were collected by flow cytometry for injection into secondary recipients. Metastasis was monitored by bioluminescence imaging in secondary recipients. All shRNAs were expressed from a pGIPZ miRNA-based construct with TurboGFP from GE Dharmacon. For ALDH1L2, the following GE Dharmacon shRNA clones were used: V2LHS_30207, V2LHS_30209. For MTHFD1 the following GE Dharmacon shRNA clones were used: V2LHS_216208 and V2LHS_196832. Mice were injected with 100 luciferase-dsRed+ cells on the right flank and monitored until tumour diameters approached 2.5 cm, at which point they were imaged along with an uninjected control mouse using an IVIS Imaging System 200 Series (Caliper Life Sciences) with Living Image software. Mice were injected intraperitoneally with 100 μl of PBS containing d-luciferin monopotassium salt (40 μg ml−1) (Biosynth) 5 min before imaging, followed by general anaesthesia 2 min before imaging. After imaging of the whole mouse, the mice were euthanized and individual organs were surgically removed and quickly imaged. The exposure time of images ranged from 10 to 60 s depending on signal intensity. The bioluminescence signal was quantified with ‘region of interest’ measurement tools in Living Image (Perkin Elmer) software. After imaging, tumours and organs were fixed in 10% neutral-buffered formalin for histopathology. For live imaging, mice were imaged once a month, and whole body bioluminescence was quantified using Living Image Software (Perkin Elmer). Mice were euthanized by cervical dislocation. Subcutaneous tumours and metastatic nodules were dissected, immediately homogenized in 80% methanol chilled with dry ice (Honeywell), vortexed vigorously, and metabolites were extracted overnight at −80 °C. The following day, samples were centrifuged at 13,000g for 15 min at 4 °C, the supernatant was collected, and metabolites from the pellet were re-extracted with 80% methanol at −80 °C for 4 h. After centrifugation, both supernatants were pooled and lyophilized using a SpeedVac (Thermo). To inhibit spontaneous oxidation, samples were extracted with 80% methanol containing 0.1% formic acid in some experiments47. Dried metabolites were reconstituted in 0.03% formic acid in water, vortexed and centrifuged, then the supernatant was analysed using liquid chromatography-tandem mass spectrometry (LC–MS/MS). A Nexera Ultra High Performance Liquid Chromatograph (UHPLC) system (Shimadzu) was used for liquid chromatography, with a Polar-RP HPLC column (150 × 2 mm, 4 μm, 80 Å, Phenomenex) and the following gradient: 0–3 min 100% mobile phase A; 3–15 min 100–0% A; 15–17 min 0% A; 17–18 min 0–100% A; 18–23 min 100% A. Mobile phase A was 0.03% formic acid in water. Mobile phase B was 0.03% formic acid in acetonitrile. The flow rate was 0.5 ml min−1 and the column temperature was 35 °C. A triple quadrupole mass spectrometer (AB Sciex QTRAP 5500) was used for metabolite detection as previously described48. Chromatogram peak areas were integrated using Multiquant (AB Sciex). To measure GSH and GSSG levels, some metabolite extractions were performed with 0.1% formic acid in 80% methanol, to inhibit spontaneous GSH oxidation. To calculate GSH and GSSG amounts, a standard curve was prepared by adding known quantities of GSH and GSSG to tumour metabolite extracts. Mice were injected intraperitoneally with 2 g kg−1 body mass of uniformly 13C-labelled glucose (Cambridge Isotopes) and were analysed 15, 30 and 60 min later. Mice were fasted for 14 h before the injection. In most experiments, subcutaneous tumours and metastatic nodules were surgically excised and homogenized in ice cold 50% methanol for GC–MS and in 80% dry ice-cold methanol for LC–MS analysis. Metabolites were extracted with three freeze-thaw cycles in liquid nitrogen. Supernatant was collected after a 15 min centrifugation at 13,000g at 4 °C and lyophilized. Metabolites were derivatized with trimethylsilyl (TMS) at 42 °C for 30 min for GC–MS analysis. 13C-enrichment analysis was performed by GC–MS as previously described48. For LC–MS analysis, lyophilized samples were resuspended in either 0.03% formic acid in water or in 5 mM ammonium acetate in water depending on the method of analysis. For 13C-enrichment analysis of lactate, serine and glycine by LC–MS/MS, we used the liquid chromatography procedure described above for LC–MS/MS metabolomics analysis with the following modifications: the liquid chromatography gradient was 0–3 min 100% mobile phase A; 3–15 min 100–0% A; 15–17 min 0% A; 17–17.5 min 0–100% A; 17.5–20 min 100% A. For analysis of 3-PG, the liquid chromatography conditions were: mobile phase A, 5 mM ammonium acetate in water and mobile phase B, 5 mM ammonium acetate in acetonitrile, and a Fusion-RP HPLC column (150 × 2 mm, 4 μm, 80 Å, Phenomenex). The liquid chromatography gradient was: 0–3 min 100% mobile phase A; 3–9 min 100–0% A; 9–11 min 0% A; 11–12 min 0–100% A; 12–15 min 100% A. For metabolite detection a triple quadrupole mass spectrometer (AB Sciex QTRAP 5500) was used on multiple reaction monitoring mode as previously described, with some modifications48. The following transitions were used: positive mode: serine 106.1/60 (M + 1: 107.1/60 and 107.1/61, M + 2: 108.1/61 and 108.1/62, M + 3: 109.1/62), glycine 76/30 (M + 1 77/30 and 77/31, M + 2 78/31); negative mode: lactate 89/43 (M + 1 90/43 and 90/44, M + 2 91/44 and 91/45, M + 3 92/45), 3-PG 185/79 (M + 1 186/79, M + 2 187/79, M + 3 188/79) and 185/97 (M + 1 186/97, M + 2 187/97, M + 3 188/97). Unlabelled tissue was used as a negative control to confirm isotopic labelling in specific transitions. Melanomas were generally dissociated enzymatically as described above. Equal numbers of dissociated cells (500,000–2,000,000) from subcutaneous tumours, Ficoll-depleted blood, or metastatic nodules were loaded with dyes to assess mitochondrial mass, mitochondrial membrane potential, and ROS levels. The dyes that were used to assess these parameters were all obtained from Life Technologies. We stained the dissociated cells for 20–45 min at 37 °C with 5 μM Mitotracker Green, Mitotracker DeepRed, CellROX Green, or CellROX DeepRed in HBSS-free (Ca2+- and Mg2+-free) to assess mitochondrial mass, mitochondrial membrane potential, mitochondrial and cytoplasmic ROS, respectively. For each indicator, staining intensity per cell was assessed by flow cytometry in live human melanoma cells (positive for human HLA and dsRed and negative for DAPI and mouse CD45/CD31/Ter119). All animal experiments were performed according to protocols approved by the Institutional Animal Care and Use Committee at the University of Texas Southwestern Medical Center (protocol 2011-0118). Unless otherwise stated, 100 freshly dissociated melanoma cells were injected subcutaneously into the right flanks of NSG mice. When tumours became palpable, in some experiments mice were injected subcutaneously with NAC (Sigma, 200 mg kg−1 day−1 in 200 μl PBS, pH 7.4) or PBS as a control. Mice were injected with their last NAC dose 10 min before being euthanized for end-point analysis. In experiments where mice received NAC via the drinking water, NAC was dissolved in PBS at 1 mg ml−1 and the water was changed every 2 days. In other experiments methotrexate (Tocris, 1.25 mg kg−1 day−1 in 100 μl PBS) was injected intraperitoneally 5 days per week. Mice that received methotrexate were simultaneously administered thymidine (Sigma, 3 mg per mouse per day in 100 μl PBS) and hypoxanthine (Sigma, 750 μg per mouse per day in 100 μl PBS) to prevent suppression of nucleotide biosynthesis. Tumour growth was monitored weekly with a caliper. Experiments were terminated when any tumour in the cohort reached 2.5 cm in size. At the end of experiments, blood was collected by cardiac puncture. Organs were analysed for micrometastases and macrometastases by bioluminescence imaging and visual inspection. Subcutaneous tumours or metastatic nodules were surgically excised as quickly as possible after euthanizing the mice then melanoma cells were mechanically dissociated and NADPH and NADP+ were measured using NADPH/NADP Glo-Assay (Promega) following the manufactures instructions. Luminescence was measured using a using a FLUOstar Omega plate reader (BMG Labtech). Values were normalized to protein concentration, measured using a bicinchoninic acid (BCA) protein assay (Thermo). Tissue lysates were prepared in Kontes tubes with disposable pestles using RIPA Buffer (Cell Signaling Technology) supplemented with phenylmethylsulfonyl fluoride (Sigma), and protease and phosphatase inhibitor cocktails (Roche). The BCA protein assay (Thermo) was used to quantify protein concentrations. Equal amounts of protein (15–30 μg) were separated on 4–20% Tris Glycine SDS gels (BioRad) and transferred to polyvinylidene difluoride membranes (BioRad). Membranes were blocked for 30 min at room temperature with 5% milk in TBS supplemented with 0.1% Tween20 (TBST) then incubated with primary antibodies overnight at 4 °C. After incubating with horseradish peroxidase conjugated secondary antibodies (Cell Signaling Technology), membranes were developed using SuperSignal West Pico or Femto chemiluminescence reagents (Thermo). Blots were stripped with 1% SDS, 25 mM glycine, pH 2, before re-probing. The following primary antibodies were used for western blot analyses: ALDH1L2 (LifeSpan Bio; LS-C178510), DHFR (LifeSpan Bio; LS-C138829), MTHFR (LifeSpan Bio; LS-C157974), SHMT1 (Cell Signaling; 12612S), SHMT2 (Cell Signaling; 12762S), MTHFD1 (ProteinTech; 10794-1-AP), MTHFD2 (ProtenTech; 12270-1-AP) and aActin (Abcam, ab8227). Tissues were fixed in 4% paraformaldehyde for 12 h at 4 °C, and then transferred to 30% sucrose for 24 h for cryoprotection. Tissues were then frozen in OCT. Sections (10 μm) were permeabilized in PBS with 0.2% Triton (PBT), three times for 5 min each, and blocked in 5% goat serum in PBT for 30 min at room temperature. Sections were then stained with primary antibodies overnight: ALDH1L2 (LS-C178510, LifeSpan Bio; 1:50) and S100 (Z0311, Dako, 1:500). The next day, sections were washed in PBS with 0.2% Triton and stained with secondary goat anti-rabbit antibody (Invitrogen) at 1:500 for 30 min in the dark at room temperature. Sections were washed with PBT with DAPI (1:1,000) and mounted for imaging. No statistical methods were used to predetermine sample size. The data in most figure panels reflect several independent experiments performed on different days using melanomas derived from several patients. Variation is always indicated using standard deviation. For analysis of statistical significance, we first tested whether there was homogeneity of variation across treatments (as required for ANOVA) using Levene’s test, or when only two conditions were compared, using the F-test. In cases where the variation significantly differed among treatments, the data were log -transformed. If the data contained zero values, 1/2 of the smallest non-zero value was added to all measurements before log transformation. If the data contained negative values, all measurements were log-modulus transformed (L(x) = sign(x) × log(|x| + 1)). In the rare cases when the transformed data continued to exhibit variation that significantly differed among treatments, we used a non-parametric Kruskal–Wallis test or a non-parametric Mann–Whitney test to assess the significance of differences among populations and treatments. Usually, variation did not significantly differ among treatments. Under those circumstances, two-tailed Student’s t-tests were used to test the significance of differences between two treatments. When more than two treatments were compared, a one-way ANOVA followed by Dunnett’s multiple comparisons tests were performed. A two-way ANOVA followed by Dunnett’s multiple comparisons tests were used in cases where more than two groups were compared with repeated measures. Hierarchical clustering was performed using Euclidean distance in Metaboanalyst49. Mouse cages were randomized between treatments in all in vivo experiments (mice within the same cage had to be part of the same treatment). No blinding was used in any experiment. In all xenograft assays we injected 4–8-week-old NSG mice, 5 mice per treatment. Both male and female mice were used. For long-term assays, we injected 10 mice per treatment to account for non-melanoma related deaths (NSG mice are susceptible to death from opportunistic infections). When mice died before the end of experiments due to opportunistic infections the data from those mice were excluded. There were only two experiments in which this occurred. In Fig. 1c, d, 0–4 mice per melanoma line were found dead owing to an opportunistic bacterial infection before termination of the experiment and were excluded from the reported results. In Fig. 2b, 0–3 mice per melanoma line were found dead owing to opportunistic infections, before the first imaging time point after transplantation. These mice were excluded from the reported results.

No statistical methods were used to predetermine sample size. The experiments were not randomized. The investigators were not blinded to allocation during experiments and outcome assessment. The E. coli SecB gene was cloned into the pET-16b vector (Novagen) containing a His -tag and a tobacco etch virus (TEV) protease cleavage site at the N terminus. Protein samples of E. coli PhoA were produced as described before20. All E. coli MBP constructs were cloned into the pET-16b vector containing a His -tag and a TEV protease cleavage site at the N terminus. The following MBP constructs were prepared in this study (residue numbers of the boundaries are in superscript): MBP1–396, mature MBP27–396, MBP29–99, MBP67–99, MBP97–164, MBP160–201, MBP198–265, MBP260–336, MBP331–396, and the MBP variants MBPG32D/I33P, MBPY283D and MBPV8G/Y283D (MBP mutants are numbered on the basis of the amino-acid sequence of the mature form of MBP). All constructs were transformed into BL21(DE3) cells. Isotopically unlabelled protein samples were produced in cells grown in Luria-Bertani (LB) medium at 37 °C in the presence of ampicillin (100 μg ml−1) to an absorbance at 600 nm (A   ) ≈ 0.8. Protein induction was induced by the addition of 0.2 mM isopropyl-β-d-1-thiogalactopyranoside (IPTG) and cells were allowed to grow for 16 h at 18 °C. Cells were harvested at A    ≈ 1.5 and resuspended in lysis buffer (50 mM Tris-HCl, 500 mM NaCl, pH 8 and 1 mM PMSF). Cells were disrupted by a high-pressure homogenizer and centrifuged at 50,000g. Proteins were purified using Ni Sepharose 6 Fast Flow resin (GE Healthcare), followed by tag removal by TEV protease at 4 °C (incubation for 16 h) and gel filtration using Superdex 75 16/60 or 200 16/60 columns (GE Healthcare). Protein concentration was determined spectrophotometrically at 280 nm using the corresponding extinction coefficient. MALS was measured using DAWN HELEOS-II (Wyatt Technology Corporation) downstream of a Shimadzu liquid chromatography system connected to a Superdex 200 10/300 GL (GE Healthcare) gel filtration column. The running buffer for SecB−PhoA complexes was 20 mM KPi (pH 7.0), 100 mM KCl, 4 mM βME, and 0.5 mM EDTA, whereas for SecB−MBP complexes was 20 mM HEPES, pH 7, 150 mM KOAc and 0.05% NaN . Protein samples at a concentration of 0.05–0.2 mM were used. The flow rate was set to 0.5 ml min−1 with an injection volume of 200 μl and the light scattering signal was collected at room temperature (~23 °C). The data were analysed with ASTRA version 6.0.5 (Wyatt Technology Corporation). ITC was performed using an iTC200 microcalorimeter (GE Healthcare) at temperatures ranging from 4 °C to 25 °C. All protein samples were extensively dialysed against the ITC buffer containing 50 mM KPi (pH 7.0), 50 mM KCl, 0.05% NaN and 2 mM tris(2-carboxyethyl)phosphine (TCEP). All solutions were filtered using membrane filters (pore size, 0.45 μm) and thoroughly degassed for 20 min before the titrations. The 40-μl injection syringe was filled with ~0.05–1 mM of SecB solution and the 200-μl cell was filled with ~0.01–0.2 mM PhoA or MBP. To measure the binding affinity of MBP to SecB, the slowly folding MBPV8G/Y283D variant was used to measure the affinity of MBP for SecB. MBPV8G/Y283D was unfolded in 8 M urea, 20 mM HEPES, pH 7, 150 mM KOAc and 0.05% NaN , and diluted 20 times to give a final concentration of 2.7 μM immediately before loading into the cell. The solution containing SecB was precisely adjusted to match the urea concentration. The titrations were performed with a preliminary 0.2-μl injection, followed typically by 15 injections of 2.5 μl each with time intervals of 3 min. The solution was stirred at 1,000 r.p.m. Data for the preliminary injection, which are affected by diffusion of the solution from and into the injection syringe during the initial equilibration period, were discarded. Binding isotherms were generated by plotting heats of reaction normalized by the modes of injectant versus the ratio of total injectant to total protein per injection. The data were fitted with Origin 7.0 (OriginLab Corporation). Isotopically labelled samples for NMR studies were prepared by growing the cells in minimal (M9) medium. Cells were typically harvested at A    ≈ 1.0. U-[2H,13C,15N]-labelled samples were prepared for the backbone assignment of SecB and large MBP fragments by supplementing the growing medium with 15NH Cl (1 g l−1) and 2H ,13C -glucose (2 g l−1) in 99.9% 2H O (CIL and Isotec). The 1H–13C methyl-labelled samples were prepared as described20, 29, 31. α-Ketobutyric acid (50 mg l−1) and α-ketoisovaleric acid (85 mg l−1) were added to the culture 1 h before the addition of IPTG. Met-[13CH ]- and Ala-[13CH ]-labelled samples were produced by supplementing the medium with [13CH ]-Met (50 mg l−1) and [2H ,13CH ]-Ala (50 mg l−1). For Thr labelling, a Thr-auxotrophic cell strain was used, and the medium was supplemented with [2H ,13CH ]-Thr (25 mg l−1). For Phe, Tyr, and Trp labelling, U-[1H,13C]-labelled amino acids were used. Alternative 13C-labelling of aromatic residues was performed as described32. All precursors and amino acids were added to the culture 1 h before the addition of IPTG, except Ala, which was added 30 min before induction. NMR samples were typically prepared in 50 mM KPi (pH 7.0), 50 mM KCl, 0.05% NaN , 5 mM βME and 7% D O. NMR experiments were recorded on Bruker 900, 850 and 700 MHz spectrometers. NMR spectra were typically recorded at 10 °C for the isolated PhoA and MBP fragments and at 35 °C for SecB and its complexes. Protein sample concentration ranged from 0.1 to 1.0 mM. All NMR spectra were processed using NMRPipe33 and analysed using NMRView (http://www.onemoonscientific.com). The SecB tetramer packs as a dimer of dimers and gives rise to two pairs of magnetically equivalent subunits: A and D give one set of resonances and subunits B and C give another set of resonances (Extended Data Fig. 1a). Sequential backbone assignment of SecB was achieved by the use of standard triple-resonance NMR pulse sequences. Three-dimensional (3D) 1H–15N NOESY experiments were used to confirm and extend the backbone assignment within each subunit. Side-chain assignment for methyls and aromatic residues was accomplished using the following NMR experiments: 3D (1H)–13C heteronuclear multiple-quantum coherence (HMQC)–NOESY-1H–13C HMQC, 13C-edited NOESY–HSQC, 13C-edited HSQC–NOESY, 15N-edited NOESY-HSQC, 3D (1H)–13C HSQC–NOESY-1H–15N HSQC, and 3D (1H)–15N HSQC–NOESY-1H–13C HSQC. We previously described the assignment strategy for unfolded PhoA20. We followed a similar strategy to assign MBP in the unfolded state by making use of several MBP fragments that remain soluble and unfolded when isolated (Extended Data Fig. 1c): MBP29–99, MBP67–99, MBP97–164, MBP160–201, MBP198–265, MBP260–336 and MBP331–396. Isolated MBP fragments encompassing the first 26 N-terminal residues (signal sequence) were not stable and this region could only be assigned in complex with SecB. Overlay of the spectra of the MBP fragments with the spectra of full-length MBP in 4 M urea indicated very good resonance correspondence. This is expected because all of the fragments, as well as the MBP, in 4 M urea are unfolded. Resonance assignment obtained for the various fragments was transferred to full length MBP in urea, and ambiguities were resolved by the use of 3D NMR spectra. It should be noted that although resonance dispersion in unliganded PhoA and MBP is poor, complex formation with SecB alleviates this problem (for the PhoA and MBP residues in the SecB-binding regions) with the spectra being of high resolution (Extended Data Fig. 4c). Assignment of the resonances in SecB−PhoA was accomplished by first assigning the complexes between SecB and the individual PhoA sites (SecB−PhoAa, SecB−PhoAc, SecB−PhoAd, SecB−PhoAe). We used U-12C,15N-labelled samples that contained specifically protonated methyl groups of Ala, Val, Leu, Met, Thr and Ile (δ1) and protonated aromatic residues Phe, Tyr and Trp in an otherwise deuterated background. The high sensitivity and resolution of the methyl region, combined with the high abundance of these nine amino acids in SecB (Extended Data Fig. 1a) and in the SecB-binding sites of PhoA and MBP, provided a large number of intermolecular NOEs for the SecB−PhoA and SecB−MBP complexes (Extended Data Table 1). Because PhoA in complex with SecB provided higher quality spectra than the spectra of MBP in complex with SecB, we determined first the structure of the SecB−PhoA complex (~120 kDa) by NMR. We initially characterized the structure of the each PhoA site (a–e) individually in complex with SecB (Extended Data Fig. 5). The structures of SecB−PhoAa, SecB−PhoAc, SecB−PhoAd, and SecB−PhoAe, were determined by NMR and are presented in Extended Data Fig. 5. A large number of intermolecular NOEs were collected for each one of the complexes (Extended Data Table 1). Because of the relatively short length of the polypeptides encompassing the individual PhoA sites, multiple PhoA molecules bound to SecB, as shown in Extended Data Fig. 5. We also note that we detected the presence of a small number of intermolecular NOEs that were suggestive of alternative conformations of the PhoA sites bound to SecB. However, the intensity of these sets of NOEs was much weaker, indicating that the population of such alternative complexes is low. To solve the structure of the SecB−PhoA complex, we sought to determine how each one of the PhoA sites binds to SecB in the context of the full length PhoA. To circumvent the signal overlap in this large complex, we used samples where the two proteins were isotopically labelled in different amino acids. For example, in one of these samples SecB was labelled in the methyls of Leu, Val and Met, whereas PhoA in the methyls of Ile amino acids. Because of the distinct chemical shifts of 1H and 13C resonances of the methyls and the isotope labelling scheme, it was possible to measure specific intermolecular NOEs between SecB and PhoA (Extended Data Fig. 4b). Several of these samples were used to determine as many intermolecular NOEs as possible. As expected, the NOEs were compatible with the structure of each PhoA site in complex with SecB, with the crucial difference that only one PhoA molecule could be accommodated in SecB. Owing to its short length, the isolated PhoA site b (PhoAb) binds to almost all of the exposed hydrophobic surface of SecB, as determined by NMR. In the SecB−PhoA complex with SecB, PhoA site b can only bind to the secondary binding site, as determined by NOEs. To further corroborate the structure of the SecB−PhoA complex we used PRE data (see below). The PRE-derived distances were fully compatible with the NOE data collected on SecB−PhoA. The structure of the SecB−PhoA complex was determined using the set of intermolecular NOEs collected directly in the complex and further refined using the intermolecular NOEs collected for the corresponding isolated PhoA sites in complex with SecB. It should be noted that because of the symmetry in SecB, the various PhoA sites may bind to any of the four SecB subunits. The final arrangement will be dictated by the length of the linkers tethering the SecB-recognition sites (as shown in Fig. 2), namely how far nearby recognition sites can bind from each other, and thus alternative routes of the polypeptide bound to SecB may be present. The only conceivable difference among the various conformations is the relative disposition of the PhoA sites. In all cases all of the SecB-recognition sites in PhoA are engaged by SecB in the complex and PhoA wraps around SecB. The NMR-driven structural model of the SecB−MBP complex (Extended Data Fig. 7b) was determined as follows: NMR analysis demonstrated that all seven recognition sites in MBP (labelled a–g) are bound to SecB in the SecB−MBP complex (Extended Data Fig. 7a). We have determined the high-resolution structure of MBPd and MBPe in complex with SecB (Extended Data Fig. 6). Because of their length and the short linker tethering the two sites, d and e sites most probably bind to the same side of SecB. MBP site f is the longest one, consisting of ~90 residues, and is thus entirely accommodated on the other side of SecB. With sites d, e and f occupying the primary binding sites, the other recognition sites (a, b, c and g), being much shorter, can be accommodated within the secondary client-binding sites on SecB. The structure of MBP sites d and e in complex with SecB was determined using the experimental intermolecular NOE data. The hydrophobic residues of the sites a, b, c, f, and g, showing the strongest effect upon SecB binding as determined by differential line broadening, were used to drive the docking of these sites to non-polar residues on SecB. The modelled structure shows that the entire MBP sequence can be accommodated within one SecB molecule. The structures of SecB in complex with PhoA and MBP were calculated with CYANA 3.97 (ref. 34), using NOE peak lists from 3D (1H)–13C HMQC–NOESY-1H–13C HMQC, 3D (1H)–15N HSQC–NOESY-1H–13C HSQC, 13C-edited NOESY–HSQC, and 15N-edited NOESY–-HSQC. The 13Cα, 13Cβ, 13C′, 15N and NH chemical shifts served as input for the TALOS+ program35 to extract dihedral angles (φ and ψ). The side chains of SecB residues within or nearby the PhoA and MBP binding sites were set flexible and their conformation was determined using intermolecular NOEs collected for each one of the complexes. The SecB regions remote to the binding sites were set rigid using the crystal structure coordinates for E. coli SecB26. The 20 lowest-energy structures were refined by restrained molecular dynamics in explicit water with CNS36. The percentage of residues falling in favoured and disallowed regions, respectively, of the Ramachandran plot is as follows: 99.4% and 0.6% for SecB–PhoA; 99.4% and 0.6% for SecB–PhoAa; 99.3% and 0.7% for SecB–PhoAc; 99.2% and 0.8% for SecB–PhoAd; 99.3% and 0.7% for SecB–PhoAe; 99.4% and 0.6% for SecB–MBPd; and 99.4% and 0.6% for SecB–MBPe. PRE experiments were used to confirm the position of each individual PhoA binding site in the SecB−PhoA complex. First, a ‘Cys-free’ variant of PhoA was prepared by mutating the four naturally occurring Cys residues in PhoA (Cys190, Cys200, Cys308 and Cys358) to Ser. We then introduced a Cys residue to either end of each SecB-binding site in PhoA and prepared a total of ten single-Cys mutants: T5C, T23C, K65C, M75C, G91C, G140C, Q274C, C308, N450C and C472. The protein purified from Ni-NTA column was quickly concentrated and loaded onto HiLoad 16/60 Superdex 200 gel filtration column (GE healthcare) using a buffer containing 50 mM KPi (pH 7.0), 150 mM NaCl and 0.05% NaN . Immediately after elution the purified single-Cys PhoA mutant was divided into two equal portions for parallel treatment with (1-oxyl-2,2,5,5-tetramethyl-3-pyrroline-3-methyl)-ethanethiosulfonate (MTSL, Toronto Research Chemicals, Toronto) and a diamagnetic MTSL analogue, in a tenfold molar excess at 4 °C for 16–20 h. MTSL was prepared in a 50 mM concentrated stock in acetonitrile. Free MTSL was removed by extensive buffer exchange using Centricon Centrifugal Filter with a MWCO of 10,000 (Millipore) at 4 °C. The MTSL-labelled PhoA protein samples were then concentrated and added into the 2H-methyl-13CH -labelled SecB at a final molar ratio of PhoA:SecB = 1:1. 2D 1H,13C HMQC spectra were recorded at 28 °C. A sample of SecB in complex with PhoA cross-linked to a diamagnetic MTSL analogue was used as a reference. Residues experienced significant NMR signal intensity reduction (>50% intensity loss) were identified as sites being within 20 Å of the paramagnetic centre whereas residues experiencing more than 90% intensity loss were identified as sites being within 14 Å of the paramagnetic centre. Refolding experiments of MBP were performed as described before37 with some modifications. Briefly, MBP was first denatured in 8 M urea, 100 mM HEPES, 20 mM KOAc, 5 mM Mg(OAc) , pH 7.4, and 0.05% NaN . Refolding was initiated by rapid dilution (20 times dilution) in the urea-free buffer and the refolding process of MBP in the absence and presence of SecB or TF was monitored by the change in the intrinsic Trp fluorescence. Fluorescence intensity was measured using either a spectrofluorometer (FluoroMax-4, Horiba) or a microplate reader (Infinite 200 PRO, Tecan). The excitation and emission wavelengths were set to 295 nm and 345 nm, respectively. For measurement using the FluoroMax-4 instrument, the MBP concentration in the 1-ml cuvette was 0.4 μM, whereas for the microplate reader experiments the concentration of MBP was 4 μM in the 30 μl-plate well. All fluorescence measurements were performed at 25 °C. Data were fitted by the Prism 6 (GraphPad) software using the nonlinear regression analysis equation38. All SPR experiments were performed on a Biacore T200 system (GE Healthcare) using a NTA-coated Sensor Chip NTA (GE Healthcare) at a flow rate of 50 μl min−1. The PhoA protein sample used for SPR experiments was genetically constructed with a His -tag at the carboxy (C) terminus and a flexible (Gly-Ser) linker repeat inserted in between to avoid steric hindrance. A single-cycle kinetic procedure was used to characterize the interaction of SecB and PhoA. The His-tagged PhoA was immobilized onto a NTA sensor chip, followed by washing with the running buffer containing 50 mM phosphate, 50 mM KCl, pH 7, 0.05% NaN , and 2 mM TCEP. The reducing agent (TCEP) ensured that PhoA was in the unfolded state20. SecB (analyte) at a range of concentrations (0.1–25.6 μM) was injected, and data for a period of 30 s of association and 60 s of dissociation were collected. MBP was prepared with a His -tag at the N terminus followed by a flexible nine-residue linker to avoid steric hindrance. Multiple-cycle kinetic analysis was performed for the SPR experiments of the binding between MBP and SecB where each sample concentration was run in a separate cycle, and the surface was regenerated between each cycle using NTA regeneration buffer. His-tagged MBP was denatured in 8 M urea and immobilized onto a NTA sensor chip. Urea was quickly washed away by running buffer containing 20 mM HEPES, pH 7.4, 150 mM KOAc and 0.05% NaN . SecB was injected at concentrations ranging from 2.5 nM to 1.6 μM. The association and dissociation time for data collection was set as 90 s and 120 s, respectively. After urea was removed, MBP remained in the unfolded conformation for sufficient time to interact with SecB. This was confirmed by monitoring the refolding behaviour of MBP using an Infinite 200 PRO microplate reader (Tecan) at the temperature range of the experiments. All SPR experiments were repeated three times and highly reproducible data were obtained. The sensorgrams obtained from the assay channel were subtracted by the buffer control, and data were fitted using the Biacore T200 evaluation software (version 1.0). BLI experiments were performed using an Octet system (forteBIO) at room temperature (~23 °C). MBP was biotinylated using the biotination kit EZ-Link NHS-PEG4-Biotin (Thermo Fisher Scientific). Biotin label freshly dissolved in water was added to the protein solution to a final molar ratio of 1:1 in buffer containing 50 mM KPi, pH 7, 150 mM NaCl, 0.05% NaN , and the solution was mixed at room temperature for 45 min. Unlabelled biotin label was removed by extensive buffer exchange using Centricon Centrifugal Filter with a MWCO of 10,000 (Millipore) at 4 °C using a buffer containing 20 mM HEPES (pH 7), 150 mM KoAc and 0.05% NaN . Biotin-labelled MBP (200 nM) denatured in 8 M urea was immobilized onto the streptavidin (SA) biosensor, and the biosensors were subsequently blocked with biocytin in 8 M urea solution before a quick 30 s dip into the urea-free buffer. SecB or TF previously diluted was applied in a dose-dependent manner to the biosensors immobilized with MBP. Bovine serum albumin (BSA) powder (Sigma-Aldrich) was added to a final concentration of 2% to avoid non-specific interaction. Parallel experiments were performed for reference sensors with no MBP captured and the signals were subsequently subtracted during data analysis. The association and dissociation periods were set to 2 min and 5 min, respectively.

As the low hanging lightweighting fruits are picked, automotive manufacturers have to now work that bit harder to shed the pounds. BMW is perhaps one of the more progressive examples of a manufacturer driving composite R&D in a major way. However, it is not simply abandoning metal, and like many others, it is seeking out ways to lightweight vehicles using conventional steel. This has led to BMW Mini initiating a project between The University of Oxford and Diamond Light Source – the UK's national synchrotron science facility located at the Harwell Science and Innovation Campus in Oxfordshire. The facility harnesses the power of electrons and x-rays to help scientists and engineers gain new insight and understanding in to the microscopic and internal structures of materials. BMW Mini wants answers to a phenomenon that has been witnessed, more broadly, since the 1950s. It is seen when parts are stamp formed, a common automotive process, to make everything from bonnets to doors. The problem is, for any stamping process that is completed in more than one stage, the deformation becomes highly complex, particularly for a pressed part with strain applied in two-axes. This results in a non-homogeneous arrangement of the crystalline and microscopic defect structures. While it may sound arbitrary, the affect can influence grain morphology, crystal orientation and distribution – all of which have significant impact on mechanical properties, including most importantly, how much the material will stretch before it fractures. Leading the work is Dr David Collins, a researcher at the University of Oxford, who explains the problem. He says: "They can't stamp form stronger steels at the moment because of this affect. The metals they use on body panels are actually quite weak, say 10% as strong as the strongest steels on the market. Stronger metals are not ductile and can't be stamped into the complicated shapes needed." It means thicker sections have to be used, so panels end up weighing more. It's something BMW is keen to understand so it can ultimately reduce the weight of steel chassis. The problem is complex, meaning testing and analysis is far from straightforward. The deformation is biaxial, meaning any obvious traditional solution is not suitable. It led to Dr Collins applying to use the synchrotron to carry out tests and shed new light, or x-rays in this case, on the problem. However, he soon faced another problem, the Shimadzu AGS-X 10kN load frame that was available, was a single axis machine. Dr Collins decided to do what so many engineers have done before him – innovate. He designed a mechanism to generate the bi-axial deformation needed, which also bolts straight on to the Shimadzu machine. "I've built a mechanism that uses the power of the load frame to generate the stresses," he says. "There are four diagonal rods at the corners to change the angle and determine the ratio of how much it is deforming in each direction. I can make it more biased towards the horizontal, vertical or even one side. "I spent time in the workshop machining all the components myself. I started from scratch and it was a very steep learning curve. I broke a lot of tools and upset a couple of people, but I was very determined to make the mechanism and run the tests." The request to use the synchrotron was successful and four days of testing commenced. The results show a series of concentric rings, with each ring corresponding to a signal coming from individual lattice planes. The shape of these rings, and their radius, gives an important microscopic insight. "If you deform a bit of material, those rings change in diameter," he says. "And you can then measure how much strain is being taken up by individual planes. So you capture what is happening on the atomic planar level." The cross shaped specimen is 1mm thick, but in the centre this is reduced to just 300µm. If it was a uniform thickness, the test would simply pull the arms off. A cross is machined in the middle, which steps down to a thinner circular cross section just 300µm thick. "One thing you can see is how strain is being accommodated in the individual grains based on their orientations," says Dr Collins. "You can monitor the centre on a macroscopic level by putting a camera in front of the rig. But with x-rays, they tell you what the strain is, on all of the individual crystals." With the tests complete, the hard work begins. After four intense days of gathering data, it could be a year or more of analysis to find conclusive explanation of the bi-axial deformation phenomenon. What is known is that 'material texture', ie the orientation of grains in the material, has a big effect on the ductility of the materials and how strain is accumulated. As all sheet materials are rolled during production, it forces most of the crystals to align in one orientation. "We can tell BMW things like 'texture is important', but we don't yet know how to optimise it," says Dr Collins. "Eventually, however, we hope to get enough of an idea about the affect to apply it to different materials. We're sticking with steel for the moment as we don't want to end up blurring the problem with other complexities. But, there is no reason why this research could not be limited to any alloy. For a lot of metals, no one has proved if this phenomenon even exists or not, so there might be massive benefits with lots of different applications outside the automotive world." Explore further: A new twist makes for better steel

With the exception of tumour xenograft studies, no statistical methods were used to predetermine sample size. The experiments were not randomized. The investigators were not blinded to allocation during experiments and outcome assessment. All solvents and reagents were used as obtained from commercial sources. Starting materials were purchased from commercial sources or prepared according to methods reported in the literature. Reactions involving air- or moisture-sensitive reagents were performed under a nitrogen atmosphere. NMR spectra were recorded in deuterated solvent with an Agilent 400 MHz MR DD2 spectrometer system equipped with an Oxford AS400 magnet. Chemical shifts are expressed as δ units and referenced to the residual 1H or 13C solvent signal. All coupling constants (J) are reported in hertz (s, singlet; d, doublet; t, triplet; q, quartet; m, multiplet; br, broad peak; dd, doublet of doublets; ddd, doublet of doublet of doublets; dm, doublet of multiplets). Mass spectra were measured with a Shimadzu LCMS-2020 spectrometer coupled to a Shimadzu 20A high-performance liquid chromatography (HPLC) system operating in reverse mode. Analytical purity was greater than 95% for final compounds and was determined using the following HPLC method. Buffer A: 5% acetonitrile, 95% water, 0.01% formic acid; buffer B: 95% CH CN, 5% water, 0.01% formic acid; SiliaChrom XDB C18, 5 μm, 2.1 × 50 mm, 5–95% B in 6 min, 1.0 ml min−1, 220 nm and 254 nm, electrospray-ionization-positive (ESI-positive), 300–800 atomic mass units. (2,6-Difluoro-3-nitrophenyl)(5-iodo-1H-pyrrolo[2,3-b]pyridin-3-yl)methanone. 5-Iodo-1H-pyrrolo[2,3-b]pyridine (160 g, 0.656 mol) and aluminium chloride (525 g, 3.94 mol) in nitromethane (1,640 ml) were allowed to stir at room temperature (20–25 °C) for 1 h. Then 2,6-difluoro-3-nitrobenzoyl chloride (218 g, 0.984 mmol) in nitromethane (1,640 ml) was added and the mixture was heated at 50 °C for 4 days. After cooling to 0 °C, the reaction was quenched with the dropwise addition of methanol (1.5 l), resulting in a precipitate. The mixture was diluted with water (2 l) and filtered. The crude product was triturated with methyl tert-butyl ether and filtered to give the title compound as a tan solid which was used directly in the next step (281 g, theory) without further purification. 1H NMR (400 MHz, dimethylsulfoxide (DMSO)-d ) 13.18 (br s, 1 H), 8.82 (s, 1 H), 8.62 (s, 1 H), 8.46 (m, 1 H), 8.40 (s, 1 H), 7.55 (m, 1 H). (3-Amino-2,6-difluorophenyl)(5-iodo-1H-pyrrolo[2,3-b]pyridin-3-yl)methanone. To (2,6-difluoro-3-nitrophenyl)(5-iodo-1H-pyrrolo[2,3-b]pyridin-3-yl)methanone (281 g, 656 mmol) in ethyl acetate (10.9 l) and tetrahydrofuran (10.9 l) was added tin(II) chloride dihydrate (517 g, 2.29 mol) portionwise while heating at 60 °C. The reaction mixture was held at this temperature overnight. After cooling to room temperature, the reaction mixture was quenched with 50% saturated aqueous sodium bicarbonate (1:1 water and saturated aqueous sodium bicarbonate) and filtered through Celite washing the cake with ethyl acetate. The layers were separated and the organic layer was washed with brine and then concentrated under reduced pressure to give the crude product, which was triturated with methyl tert-butyl ether and filtered to give the title compound as a tan solid (216 g, 541 mmol, 83% yield). 1H NMR (400 MHz, DMSO-d ) 12.96 (br s, 1 H), 8.72 (s, 1 H), 8.56 (d, J = 2.0 Hz, 1 H), 8.06 (s, 1 H), 6.92 (dd, J = 8.6 Hz, 1 H), 6.88 (m, 1 H), 5.20 (s, 2 H); liquid chromatography–mass spectrometry (LC/MS) (ESI-positive) m/z: 399.9 (M + H+). (3-Amino-2,6-difluorophenyl)(5-(2-cyclopropylpyrimidin-5-yl)-1H-pyrrolo[2,3-b]pyridin-3-yl)methanone. A mixture of (3-amino-2,6-difluorophenyl)(5-iodo-1H-pyrrolo[2,3-b]pyridin-3-yl)methanone (93 g, 233 mmol), 2-cyclopropyl-5-(4,4,5,5-tetramethyl-1,3,2-dioxaborolan-2-yl)pyrimidine (229 g, 466 mmol, ~ 50% purity), potassium carbonate (97.0 g, 702 mmol), and [1,1′-bis(diphenylphosphino)ferrocene]dichloropalladium(II) dichloromethane complex (19.0 g, 23.3 mmol) in dioxane (930 ml) and water (465 ml) was heated at 100 °C for several hours. Upon cooling, the reaction mixture was diluted with water and extracted with a mixture of tetrahydrofuran and ethyl acetate. The organic layer was separated and concentrated under reduced pressure to give the crude product, which was triturated with dichloromethane/methyl tert-butyl ether and filtered, washing with methyl tert-butyl ether to give the title compound as a tan solid (71.0 g, 78% yield). 1H NMR (400 MHz, DMSO-d ) 12.95 (br s, 1 H), 9.07 (s, 2 H), 8.71 (d, J = 2.3 Hz, 1 H), 8.66 (s, 1 H), 8.11 (s, 1 H), 6.92 (dd, J = 9.0 Hz, 9.0 Hz, 1 H), 6.89 (ddd, J = 5.9 Hz, 9.0 Hz, 9.0 Hz, 1 H), 5.20 (s, 2 H), 2.27 (m, 1 H), 1.03–1.22 (m, 4 H); LC/MS (ESI-positive) m/z: 392.2 (M + H+). 5-(2-Cyclopropylpyrimidin-5-yl)-3-[3-[[ethyl(methyl)sulfamoyl]amino]-2,6-difluoro-benzoyl]-1H-pyrrolo[2,3-b]pyridine (PLX7904). To (3-amino-2,6-difluorophenyl)(5-(2-cyclopropylpyrimidin-5-yl)-1H-pyrrolo[2,3-b]pyridin-3-yl)methanone (53.8 g, 138 mmol) in pyridine (1375 ml) was added ethyl(methyl)sulfamoyl chloride (65.0 g, 412 mmol) and the reaction was heated at 65 °C overnight. The volatiles were removed under reduced pressure and the residue was partitioned between water and ethyl acetate/tetrahydrofuran. The organic layer was concentrated under reduced pressure to give the crude product, which was dry loaded onto silica gel and purified by silica gel column chromatography (twice) eluting with 0–10% methanol/dichloromethane, then purified by silica gel column chromatography eluting with ethyl acetate. The fractions containing the desired product were pooled and concentrated under reduced pressure. The resulting solid was triturated with methyl tert-butyl ether and filtered to give the title compound as a white solid (21.1 g, 30% yield). 1H NMR (400 MHz, DMSO-d ) 13.07 (br s, 1 H), 9.71 (br s, 1 H), 9.03 (s, 2 H), 8.76 (s, 1 H), 8.68 (s, 1 H), 8.19 (s, 1 H), 7.59 (ddd, J = 5.9 Hz, 9.0 Hz, 9.0 Hz, 1 H), 7.27 (dd, J = 9.0 Hz, 9.0 Hz, 1 H), 3.12 (q, J = 7.0 Hz, 2 H), 2.74 (s, 3 H), 2.29 (m, 1 H), 1.03–1.22 (m, 4 H), 0.95 (t, J = 7.0 Hz, 3 H); 13C NMR (100 M Hz, DMSO-d6) 181.1, 170.4, 156.1 (dd, J = 246 Hz, J = 6.9 Hz), 155.5, 152.4 (dd, J = 250 Hz, J = 8.4 Hz), 149.7, 144.3, 139.2, 128.9, 128.6 (d, J = 9.9 Hz), 127.7, 126.2, 123.0 (dd, J = 13.3 Hz, J = 3.4 Hz), 118.5 (dd, J = 24.6 Hz, J = 22.5 Hz), 117.9, 116.2, 112.7 (dd, J = 22.5 Hz, J = 3.4 Hz), 45.3, 34.4, 18.2, 13.3, 10.9; LC/MS (ESI-positive) m/z: 513.3 (M + H+). (3R)-N-[3-[5-(2-cyclopropylpyrimidin-5-yl)-1H-pyrrolo[2,3-b]pyridine-3-carbonyl]-2,4-difluoro-phenyl]-3-fluoro-pyrrolidine-1-sulfonamide (PLX8394). This material was prepared in a manner analogous to PLX7904 using (3R)-3-fluoropyrrolidine-1-sulfonyl chloride in place of ethyl(methyl)sulfamoyl chloride. The product was purified by reverse-phase HPLC to provide, after lyophilization, the title compound as a white solid. 1H NMR (400 MHz, DMSO-d6) 13.05 (br s, 1 H), 9.84 (br s, 1 H), 9.01 (s, 2 H), 8.73 (s, 1 H), 8.67 (s, 1 H), 8.15 (s, 1 H), 7.62 (ddd, J = 5.9 Hz, 9.0 Hz, 9.0 Hz, 1 H), 7.26 (dd, J = 9.0 Hz, 9.0 Hz, 1 H), 5.29 (dm, J = 51.6 Hz (H-F), 1 H), 3.43 (dm, 2 H), 3.33 (m, 2 H), 2.27 (m, 1 H), 2.04 (m, 2 H), 1.01–1.11 (m, 4 H); 13C NMR (100 MHz, DMSO-d6) 181.1, 170.4, 156.2 (dd, J = 247 Hz, J = 6.9 Hz), 155.5, 152.6 (dd, J = 249 Hz, J = 8.4 Hz), 149.7, 144.3, 139.2, 128.9, 128.7 (d, J = 9.2 Hz), 127.7, 126.2, 122.9 (dd, J = 13.7 Hz, J = 3.8 Hz), 118.5 (dd, J = 24.4 Hz, J = 22.2 Hz), 117.9, 116.2, 112.7 (dd, J = 22.9 Hz, J = 3.9 Hz), 93.4 (d, J = 175 Hz), 54.9 (d, J = 22.9 Hz), 46.5, 32.5 (d, J = 21.3 Hz), 18.2, 10.9; LC/MS (ESI-positive) m/z: 542.9 (M + H+). 2-Tert-butyl-5-(2-chloropyrimidin-4-yl)-4-[3-[[ethyl (methyl)sulfamoyl]amino]-2-fluoro-phenyl]thiazole. To a solution of 3-[2-tert-butyl-5-(2-chloropyrimidin-4-yl) thiazol-4-yl]-2-fluoroaniline (102 mg, 0.281 mmol) in dichloromethane (1 ml) was added pyridine (0.5 ml) followed by ethyl(methyl)sulfamoyl chloride (265 mg, 1.68 mmol). The reaction was allowed to stir at 50 °C for 96 h. The reaction was worked up by extraction with ethyl acetate and 0.1 M HCl (aq). The product was purified by flash chromatography (5–30% ethyl acetate in hexanes) which gave impure material. This material was again purified by flash chromatography (0.5–6% methanol in dichloromethane). This provided the title compound (55 mg, 41% yield), which was used in the next step. 1H NMR (400 MHz, CD CN) 8.45 (d, J = 5.4 Hz, 1 H), 7.66 (t, J = 7.5 Hz, 1 H), 7.55 (s, 1 H), 7.38 (t, J = 7.5 Hz, 1 H), 7.34 (dd, J = 8.0 Hz, 1 H), 7.04 (d, J = 5.4, 1 H), 3.19 (q, J = 7.2 Hz, 2 H), 2.79 (s, 3H), 1.51 (s, 9 H), 1.09 (t, J = 7.3 Hz, 3 H); LC/MS (ESI-positive) m/z: 484.2 (M + H+). 5-(2-Aminopyrimidin-4-yl)-2-tert-butyl-4-[3-[[ethyl(methyl)sulfamoyl]amino]-2-fluoro-phenyl]thiazole (PLX7922). A solution of 2-tert-butyl-5-(2-chloropyrimidin-4-yl)-4-[3-[[ethyl(methyl)sulfamoyl]amino]-2-fluoro-phenyl]thiazole (51 mg, 0.11 mmol) dissolved in 5 ml of 7 M ammonia in methanol in a sealed reaction vial was placed in an oil bath at 80 °C and allowed to stir. After 48 h, the reaction was concentrated under reduced pressure and the resulting residue was purified by reverse-phase HPLC to provide the title compound, after lyophilization, as a white solid (31 mg, 61% yield). 1H NMR (400 MHz, DMSO-d6) 9.71 (br s, 1 H), 8.04 (d, J = 5.1 Hz, 1 H), 7.54 (m, 1 H), 7.30 (m, 2 H), 6.77 (br s, 2 H), 6.03 (d, J = 5.1 Hz, 1 H), 3.06 (q, J = 7.0 Hz, 2 H), 2.67 (s, 3 H), 1.41 (s, 9 H), 0.99 (t, J = 7.0 Hz, 3 H); 13C NMR (100 MHz, DMSO-d6) 181.9, 163.9, 159.3, 158.1, 152.2 (d, J = 251 Hz), 145.9, 134.7, 127.8, 126.9 (d, J = 13 Hz), 126.5, 125.2 (d, J = 5 Hz), 124.3 (d, J = 14 Hz), 105.8, 45.3, 38.1, 34.5, 30.8, 13.2; LC/MS (ESI-positive) m/z: 465.2 (M + H+). N-[3-[(5-chloro-1H-pyrrolo[2,3-b]pyridin-3-yl)-hydroxy-methyl]-2,4-difluoro-phenyl]-4-(trifluoromethyl)benzenesulfonamide. To a solution of N-(2,4-difluoro-3-formyl-phenyl)-4-(trifluoromethyl)benzenesulfonamide (83.4 g, 0.228 mol) and 5-chloro-1H-pyrrolo[2,3-b]pyridine (34.8 g, 0.228 mol) in anhydrous methanol (350 ml) was added potassium hydroxide (38.4 g, 0.684 mol). The reaction mixture was stirred at room temperature, under nitrogen, for 3 h and poured into water (1 l). The product was extracted with ethyl acetate (2 × 800 ml). The organic layers were combined, washed with brine (800 ml), dried, and concentrated under reduced pressure to yield a brown solid. This solid was suspended in acetonitrile (10 vol) overnight with stirring and then cooled in an ice bath for 3 h. The solids were isolated by filtration, washed with a minimum of cold acetonitrile, and dried to provide the title compound (56.8 g, 48% yield). 1H NMR (400 MHz, DMSO-d6) 11.77 (s, 1 H), 10.38 (s, 1 H), 8.17 (d, J = 2.3 Hz, 1 H), 7.88 (s, 4 H), 7.75 (d, J = 2.3 Hz, 1 H), 7.18 (s, 1 H), 7.17 (m, 1 H), 7.05 (t, J = 9.0 Hz, 1 H), 6.20 (d, J = 4.9 Hz, 1 H), 6.02 (d, J = 4.9 Hz, 1 H); (LC/MS (ESI-positive) m/z: 518.0 (M + H+). N-[3-(5-chloro-1H-pyrrolo[2,3-b]pyridine-3-carbonyl)-2,4-difluoro-phenyl]-4-(trifluoromethyl)benzenesulfonamide (PLX5568). To a solution of N-[3-[(5-chloro-1H-pyrrolo[2,3-b]pyridin-3-yl)-hydroxy-methyl]-2,4-difluoro-phenyl]-4-(trifluoromethyl)benzenesulfonamide (100 g, 0.193 mol) in tetrahydrofuran (2.5 l) was added to Dess–Martin periodinane (99.2 g, 0.234 mol) under nitrogen. When the reaction was complete, the mixture was poured into 1 M sodium thiosulfate (700 ml) and saturated sodium bicarbonate solution (700 ml) and then extracted with ethyl acetate (2 × 700 ml). The organic layers were combined, washed with brine (800 ml), dried, and concentrated under reduced pressure to yield a brown solid. This residue was stirred in ethyl acetate (1 l) and silica (100 g) for 45 min and diluted with hexane (500 ml). The mixture was poured through a plug of silica and the product was eluted with 50:50 hexane:ethyl acetate. The fractions containing the product were combined and concentrated under reduced pressure to yield crude product as a yellow solid. Recrystallization of the crude product from ethanol provided the title compound as a pale yellow solid (71 g, 71% yield). 1H NMR (400 MHz, DMSO-d6) 13.13 (s, 1 H), 10.51 (s, 1 H), 8.43 (s, 1 H), 8.38 (d, J = 2.4 Hz, 1 H), 8.18 (d, J = 2.4 Hz, 1 H), 7.93 (s, 4 H), 7.44 (m, 1 H), 7.28 (m, 1 H); 13C NMR (100 MHz, DMSO-d6) 180.7, 157.0 (dd, J = 247 Hz, J = 7.3 Hz), 153.5 (dd, J = 251 Hz, J = 8.4 Hz), 148.1, 144.0, 143.9, 139.9, 133.1 (q, J = 32.3 Hz), 130.4 (d, J = 9.2 Hz), 128.7, 128.1, 126.9 (q, J = 3.8 Hz), 126.3, 123.8 (q, J = 273 Hz), 121.3 (dd, J = 13.8 Hz, J = 3.8 Hz), 118.6, 118.4 (dd, J = 25.4 Hz, J = 22.6 Hz), 115.3, 113.1 (dd, J = 23.1 Hz, J = 3.8 Hz); (LC/MS (ESI-positive) m/z: 516.1 (M + H+). Biochemical assays and kinome selectivity profiling. The in vitro RAF kinase activities were determined by measuring phosphorylation of a biotinylated substrate peptide as described previously25. PLX7904 was also tested against a panel of 287 kinases at concentrations of 1 μM in duplicate. Kinases inhibited by over 50% were followed up by IC determination. The 287 kinases represent all major branches of the kinome phylogenetic tree. The inhibition screen of 287 kinases was performed under contract as complementary panels at Invitrogen (Life Technologies) SelectScreen profiling service, DiscoverX KINOMEScan service, and Reaction Biology Corporation Kinase HotSpot service. Cell culture experiments. The B9 cell line was a gift from A. Balmain. The SK-MEL-239 and SK-MEL-239-C3 cell lines were provided by D. Solit and N. Rosen. The IPC-298 cell line was purchased from DSMZ. All other cell lines (A375, A431, COLO829, HCT116, and SKBR3) were purchased from ATCC. All cell lines were authenticated at the source by STR profiling and tested negative for mycoplasma contamination before use. Compounds dilutions were done in 100% DMSO and these titrations were diluted 500-fold in culture medium when added to cells, resulting in a final 0.2% DMSO concentration. Final compound concentrations are listed in text and figures. Phospho-ERK AlphaScreen assay. To determine the effects of compound treatment on phosphorylation of ERK1/2, cells were plated in a 96-well plate and treated with an eight-point titration of compound for 1 h at 37 °C before lysis. To detect pERK, cell lysates were incubated with streptavidin-coated AlphaScreen donor beads, anti-mouse IgG AlphaScreen acceptor beads, a biotinylated anti-ERK1/2 rabbit antibody, and a mouse antibody that recognized ERK1/2 only when it was phosphorylated on Thr202 and Tyr204. The biotinylated ERK1/2 antibody bound both to the streptavidin-coated AlphaScreen donor beads and to ERK1/2 (regardless of its phosphorylation state), and the phospho-ERK1/2 antibody bound to the acceptor beads and to ERK1/2 that was phosphorylated at Thr202/Tyr204. An increase in ERK1/2 phosphorylation at Thr202/Tyr204 brought the donor and acceptor AlphaScreen beads into close proximity, generating a signal that could be quantified on an EnVision reader (Perkin Elmer). Inhibition of ERK phosphorylation resulted in a loss of signal compared with DMSO controls. Phospho-ERK immunoblot analysis. Western blots were performed by standard techniques and analysed on an Odyssey Infrared Scanner (Li-COR Biosciences). The following antibodies were used: pERK1/2 (T202/Y204) and ERK1/2 (Cell Signaling). Growth inhibition assay. Cells were plated into a 96-well plate at a density of 3,000 cells per well and allowed to adhere overnight. Compounds were dissolved in DMSO, diluted threefold to create an eight-point titration, and added to cells. After incubation for 72 h, cell viability was examined using CellTiter-Glo (Promega). Anchorage-independent growth assay. Twenty-five thousand B9 cells were plated in each well of a six-well plate with a bottom layer of 1% and a top layer of 0.4% low melting agar (Sigma A4018) containing RPMI1640 medium with 10% FBS. For the RAF inhibitor study, B9 cells grown in soft agar were treated with vemurafenib, PLX4720 or PLX7904 at the indicated concentrations, or DMSO at 0.2% final concentration for 3 weeks. For the EGFR ligand study, B9 cells grown in soft agar were treated with AREG (R&D Systems 989-AR), TGF-α (R&D Systems 239-A), or HB-EGF (R&D Systems 259-HE) at the indicated concentrations for 3 weeks. For the vemurafinib and erlotinib combination study, B9 cells grown in soft agar were treated with vemurafenib, erlotinib, or a combination of the two compounds at the indicated concentrations, or DMSO for 3 weeks. Anchorage-independent colonies ≥ 100 μm were scored using AxioVision Rel 4.8 software (Carl Zeiss). ELISA for detecting EGFR ligands. Twenty thousand B9 cells were plated in each well of a 96-well plate and treated with DMSO control or compounds at the indicated concentrations for 48 h. Cell supernatants were collected and cells were lysed using 1 × cell lysis buffer (CST 9803). The amounts of AREG, TGF-α, and HB-EGF in cell supernatants or cell lysates were determined with the use of ELISA Development kits (R&D Systems DY989, DY239, and 259-HE-050N) according to the manufacturer’s instructions. EGFR signalling assay. B9 cells were treated with 1 μM or 5 μM vemurafinib or control vehicle for the indicated times in the absence of serum. Supernatants from treated B9 cells were then collected and added to newly plated, serum-starved (overnight) B9 cells for 10 min. Cells were washed with 1 × PBS twice, lysed, and subjected to western blot analysis. pEGFR Y1068, EGFR, pAkt S473, and Akt antibodies were purchased from Cell Signaling Technology. RAF dimerization assays. BRAF–CRAF heterodimerization was characterized in cell lysates. Cells were plated on 96-well dishes and allowed to adhere overnight at 37 °C. Cells were treated with compound or DMSO for 1 h at 37 °C before lysis in RIPA buffer containing protease and phosphatase inhibitors. The lysates were transferred to ELISA plates coated with a monoclonal CRAF capture antibody, and incubated overnight at 4 °C. Further incubations with a polyclonal BRAF detection antibody and a horseradish-peroxidase-labelled secondary antibody were done at room temperature. After incubation with TMB substrate and sulfuric acid, the signal was analysed by measuring absorbance at 450 nm on a Tecan Safire plate reader. Microarray gene expression analysis. B9 cells were plated in 1 µM vemurafenib, 1 µM PLX7904 or 0.2% DMSO vehicle control and incubated for 17 h. Cells were harvested, total RNA was isolated (RNeasy Mini Kit, Qiagen), and gene expression was measured using Affymetrix Mouse420_2 chips following the manufacturer’s instructions. Vemurafenib response genes were identified by requiring the ratio between the treated and vehicle control samples be more than 1.9 (upregulated) or less than 0.54 (downregulated). Tumour xenograft studies. All animal studies were conducted in accordance with the Institute for Laboratory Animal Research Guide for the Care and Use of Laboratory Animals and the US Department of Agriculture’s Animal Welfare Act and approved by the institutional review board at testing facilities. Sample size (number of mice per group) was selected to provide at least 80% power to detect a two s.d. difference of mean tumour volume between two groups with two-sided type I error = 1%. The same formulation was used for both COLO205 and B9 xenograft studies. The powder of the test compound was dissolved in pure N-methyl-2-pyrrolidone. Diluent consisted of PEG400:TPGS:Poloxamer 407:water (40:5:5:50). Before gavage administration, fresh stock of N-methyl-2-pyrrolidone compound solution (or N-methyl-2-pyrrolidone for vehicle) was thoroughly mixed with the diluents to make a uniform suspension. Dosing volume was 5 µl g−1. On the last day of the efficacy study, blood samples were collected at 0, 2, 4, and 8 h after last dosing, two animals per time point, for pharmacokinetic analysis. Animals were fed a standard rodent diet and water was supplied ad libitum. Tumour measurements were taken with an electronic microcalliper three times weekly. In addition, body weights were recorded at these times. Test facility investigators were blinded to the group allocation during the experiment. COLO205 tumour cells were cultured in DMEM 10% FBS 1% penicillin/streptomycin supplemented with bovine insulin, at 37 °C. Balb/C nude mice, female, 6–8 weeks old, weighing approximately 18–22 g, were inoculated subcutaneously at the right flank with COLO205 tumour cells (5 × 106) in 0.1 ml of PBS mixed with matrigel (50:50) for tumour development. The treatment was started when mean tumour size reached approximately 100 mm3, with eight mice in each treatment group randomized to balance the average weight and tumour size. B9 cells were expanded in DMEM 10% FBS 1% penicillin/streptomycin. Upon trypsinization the cells were washed three times with 20 ml RPMI, and after the final centrifugation were re-suspended, counted, and adjusted by volume to a final concentration of 5 × 107 cells per millilitre. B9 xenografts were started by injection of 5 × 106 cells subcutaneously in 6- to 7-week-old female nude Balb/c mice. Compound dosing started when the average size of tumours reached 50–70 mm3. Animals were equally distributed over treatment groups (n = 10) to balance the average tumour size and body weight. Animals were dosed orally for days 1–14 twice daily and days 15–28 once daily with vehicle, vemurafenib 50 mg per kg, or PLX7904 50 mg per kg. 12-O-tetradecanoylphorbol-13-acetate (TPA) was put on the skin of all mice twice a week during weeks 3 and 4 at a dose of 2 µg in 200 µl acetone. Expression and purification of BRAF and BRAFV600E were performed as previously described13, 25. Crystallization drops were prepared by mixing the protein solution with 1 mM of compound and the same amount of reservoir, and drops were incubated by vapour diffusion (sitting drops) at 4 °C. The mother liquor used to obtain co-crystals of PLX7904, dabrafenib, and PLX7922 with BRAFV600E consisted of 0.1 M BisTris at PH 6.0, 12.5% 2,5-hexanediol, 12% PEG3350; the reservoir used to obtain co-crystals of PLX5568 with BRAFWT contained 0.1 M MES at pH 6.0, 35% (v/v) 2-methyl-2,4- pentanediol, and 0.2 M Li SO . All co-crystals were flash-frozen with liquid nitrogen, but BRAFV600E co-crystals were soaked in a solution containing the mother liquor plus 20% glycerol, before flash-freezing. X-ray diffraction data were collected at beamline 8.3.1 at the Advanced Light Source (Lawrence Berkeley Laboratory) and beamline 9.1 at Stanford Synchrotron Radiation Lightsource (Stanford University). Data were processed and scaled using MOSFLM31 and SCALA in the CCP4 package32. All co-structures were solved using molecular replacement with the program MOLREP33. The starting models used were the inhibitor-bound BRAFV600E and BRAFWT, respectively (Protein Data Bank accession numbers 4FK3 and 1UWJ). The final models were obtained after several rounds of manual rebuilding and refinement with PHENIX34 and REFMAC35. A summary of the crystallography statistics is included in Extended Data Table 1.

No statistical methods were used to predetermine sample size. The experiments were not randomized. MB5746 is an antibiotic-sensitized E. coli strain harbouring an envA1 mutation and tolC deletion42. Consequently, MB5746 is outer membrane hyper-permeable and efflux deficient. The primary screen was performed in 1,536-well plate format. MB5746 (1:10,000 dilution of an overnight culture 2–4 × 109 CFU ml−1) was grown in cation-adjusted Mueller Hinton broth (CAMHB; BD BBL, cat. no. 212322), with or without 10 μM riboflavin (riboflavin; Sigma, cat. no. 4500-100G) supplementation. The final medium volume in each well was 5 μl, to which 50 nl DMSO containing twofold-titrated compound was transferred. Hit compounds whose activity was suppressed by riboflavin were retested with CAMH agar containing MB5746 (1:1,000 dilution of overnight culture). MICs were determined by the broth microdilution method as recommended by the Clinical and Laboratory Standards Institute with one exception: bacterial strains were tested in M9 broth. MB5746 was grown to late-exponential phase in CAMHB and spread on CAMH agar plates (BD BBL cat. no. 211438) containing twofold escalating agar MIC levels of ribocil. To establish the number of viable cells in the starting inoculum, the culture was serially diluted and plated on CAMH agar plates lacking ribocil. Resistant isolates were re-streaked on plates containing the same ribocil concentration. The frequency of resistance (FOR) was determined, dividing the number of resistant isolates by the viable CFU in the late-exponential inoculum. ribA and ribB were knocked out in strain MB5746 by λ-Red recombineering using the linear PCR product generated by amplification from pKD4 (ref. 2) with the primer pairs P1/P2 or P3/P4, respectively. Transformation and selection on kanamycin were performed as described previously43, except that 500 µg ml−1 riboflavin was added to the selection plate to maintain the auxotrophic mutants. Upon obtaining strains MB5746ΔribA::kan and MB5746 ΔribB::kan, riboflavin growth assays were performed to determine the minimal concentration of riboflavin required to maintain growth of the auxotrophic mutant. Liquid cultures of CAMHB inoculated with 1 × 105 CFU ml−1 of the mutants grown overnight on solid media were supplemented with a twofold dilution series of riboflavin ranging from 250 µg ml−1 to 0.25 µg ml−1. After 24 h of incubation at 37 °C growth was scored visually and it was determined that as little as 0.5 µg ml−1 was sufficient to maintain some growth of the auxotrophic mutants, but optimal growth was observed with a minimum of 4 µg ml−1 riboflavin. DNA templates for in vitro transcription of aptamer RNA were prepared by PCR from the pCDF-EcFMN–GFP reporter plasmid using a forward primer ApT7 (TAATACGACTCATTATAGGgcttattctcagggcg) incorporating the T7 promoter and a reverse primer ApRev (cgttactctctcccatccg). Uppercase represents additional sequences added in the primer including the T7 promoter, lowercase represents the riboswitch sequence. In vitro RNA aptamer transcription was carried out using the RiboMAX large scale RNA production system kit (P1300, Promega) using the protocols provided by the manufacturer. After extraction with phenol/cholorform the RNA aptamers were further purified by column chromatography on NAP-10 sephadex columns (GE Healthcare) and isopropanol precipitation. E. coli MB5746 and deletion mutants (ΔribA, ΔribB) were reconstituted in 10 ml trypticase soy broth (TSB, Corning, cat. no. 46-060-CI). The mutant strains were supplemented with 2.5 µg ml−1 riboflavin (Sigma, cat. no. R4500-100G) and incubated at 35 °C for 6 h with shaking at 250 r.p.m. The respective 6-h cultures were used to seed, at a ratio of 1:50 ml, TSB either with or without 2.5 µg ml−1 riboflavin in a 250 ml flask and were incubated at 35 °C for 16 h with shaking at 250 r.p.m. Overnight cultures were centrifuged at 5,000 r.p.m. for 12 min at 5 °C. Supernatant was decanted and pellets were re-suspended in 50 ml fresh TSB to remove excess riboflavin. Six tenfold serial dilutions were made in TSB from these stock cultures (1.2 × 1010 CFU ml−1 for wild type, 5.7 × 109 CFU ml−1 for ΔribA, 3.6 × 109 CFU ml−1 for ΔribB). Select serial dilutions were further diluted 1:10 into 3% gastric hog mucin for intra-peritoneal (i.p.) injection into mice. The initial dilutions without mucin were plated for quantification on TSA II (5% sheep’s blood) agar plates (BD BBL, cat. no. 221261) for the wild-type strain or 10 µg ml−1 riboflavin-infused Muller Hinton II Agar (BD BBL, cat. no. 211438) plates for mutant strains. Eleven-week-old female DBA2/J mice (Jackson Labs) were chosen for this study based on weight (~20 g) and combined into one pool. Animals were then randomly selected from this pool and placed in groups of five in separate boxes. Subjects were treated with 150 mg kg−1 i.p. cyclophosphamide (Baxter, NDC# 10019-955-50) on day −4 and 100 mg kg−1 on day −1. On day 0, five mice per group were injected i.p. with 0.5 ml of a respective dilution of bacteria in 3% mucin (6.0 × 106, 105, 104 CFU ml−1 for wild type, 2.85 × 107, 106, 105 CFU ml−1 for ΔribA, 1.8 × 107, 106, 105 CFU ml−1 for ΔribB). On day 1, subjects were euthanized via CO asphyxiation and spleens were aseptically removed, weighed and homogenized in 1.5 ml of sterile saline (Hyclone, cat. no. SH30028.03) with 10% glycerol (Fisher Scientific, cat. no. BP229-1). Tissue homogenates were serially diluted tenfold in sterile saline and selected concentrations were plated on either TSA II (5% sheep’s blood) agar plates for the wild type or MH riboflavin infused agar plates for ΔribA and ΔribB mutants. Plates were incubated at 35 °C for 24 h and CFU per g of spleen tissue were determined. No data was excluded from this study and investigator blinding was not implemented during this study. This study was approved and was in compliance with the ethical regulations set forth by the Institutional Animal Care and Use Committee (IACUC) at Merck Research Laboratories, Kenilworth, New Jersey. E. coli MB5746 was reconstituted in 10 ml trypticase soy broth (TSB, Corning, cat. no. 46-060-CI) and incubated at 35 °C for 6 h with shaking at 250 r.p.m. The 6 h culture was used to seed, at a ratio of 1:50 ml, TSB in a 250 ml flask and were incubated at 35 °C for 16 h with shaking at 250 r.p.m. The overnight culture was centrifuged at 5,000 r.p.m. for 12 min at 5 °C. Supernatant was decanted and the pellet was re-suspended in 50 ml fresh TSB to remove excess riboflavin. Nine tenfold serial dilutions were made in TSB from the culture (1.0 × 1010 CFU ml−1). The third dilution (1.0 × 107 CFU ml−1) was further diluted into 3% gastric hog mucin for i.p. injection into mice. The initial dilutions without mucin were plated for quantification on TSA II (5% sheep’s blood) agar plates (BD BBL, cat. no. 221261). Twelve-week-old female DBA2/J mice (Charles River Laboratory) were chosen for this study based on weight (~20 g) and combined into one pool. Animals were then randomly selected from this pool and placed in groups of five in separate boxes. Subjects were treated by intraperitoneal (i.p.) injection with 150 mg kg−1 of cyclophosphamide (Baxter, NDC# 10019-955-50) on day −4 and 100 mg kg−1 on day −1. On day 0, mice were inoculated i.p. with 0.5 ml of bacteria in 3% mucin (5.0 × 104 CFU ml−1; Fig. 4) or a higher inoculum of 5.0 × 105 CFU ml−1 (Extended Data Figure 8). Thirty minutes post-inoculation, mice (n = 5 per group) were treated by subcutaneous (s.c.) injection three times over 24 h with either ciprofloxacin (0.5 mg kg−1, Sigma Aldrich, cat. no. 17850-5G-F, ribocil-C (at either 120, 60, or 30 mg kg−1)) or 10% DMSO (Sigma Aldrich, cat. no. 276855-1L) sham. On day 1, subjects were euthanized via CO asphyxiation and spleens were aseptically removed, weighed and homogenized in 1.5 ml of sterile saline (Hyclone, cat. no. SH30028.03) with 10% glycerol (Fisher Scientific, cat. no. BP229-1). Tissue homogenates were serially diluted tenfold in sterile saline and selected concentrations were plated on TSA II (5% sheep’s blood) agar plates. Plates were incubated at 35 °C for 24 h and CFU per g of spleen tissue were determined. A normality test was performed to verify normal distribution of data before determining statistical significance via the one-way Bonferroni ANOVA. No data was excluded from these studies and investigator blinding was not implemented during this study. This study was approved and was in compliance with the ethical regulations set forth by the Institutional Animal Care and Use Committee (IACUC) at Merck Research Laboratories, Kenilworth, New Jersey. Overnight cultures of MB5746 or MB5746 RibocilR cells were diluted 1:50 in CAMHB and distributed (1.25 ml) into 10-ml culture tubes containing diluted ribocil (twofold dilution series) or DMSO (1%) as mock control. The treated cultures were incubated with shaking at 37 °C for about 20 h, after which the OD of the culture was determined and 500 μl was moved to a 96-well deep-well plate. After centrifugation (4,000 r.p.m.) for 10 min, the bacterial cell pellets were rinsed with lysozyme dilution buffer (10 mM Tris HCl (pH 8.0), 25 mM NaCl, 1 mM EDTA) and centrifuged again. Cell pellets were then re-suspended in 100 μl of lysozyme solution (10 mg ml−1 lysozyme (Sigma) in lysozyme dilution buffer), incubated at 37 °C for 30 min, and then frozen at −20 °C. Riboflavin, FMN and FAD concentrations in the bacterial lysates were determined using the Vitamin B2 HPLC detection kit (ImmuChrom, GmbH) and Vitamin B2 column (IC2300rp, ImmuChrom GmbH) following the procedure recommended by the manufacturer scaled for a 50 μl sample (bacterial lysate). A Shimadzu HPLC system with fluorescence detector was used at a flow rate of 1.0 ml min−1 and flavin detection was carried out at 450 nm. Flavin levels were determined for an equivalent number of cells by correcting raw AUCs using the OD ratio of the treated versus the untreated cultures. P1:tatggcaaaataagccaatacagaaccagcattatctggagaatttcatggtgcaggctggagctgcttc; P2:aagcaaatgaattacacaatgcaagagggttatttgttcagcaaatggcccatatgaatatcctccttag; P3:gactgccctgattctggtaaccataattttagtgaggtttttttaccatggtgcaggctggagctgcttc; P4:gattaaggcagtaaattaagcagcggttttcagctggctttacgctcatgcatatgaatatcctccttag; P5:CTCAAATGCCTGAGGTTTCAGcaggacttgcgtttggacgtc; P6:GAAAAGTTCTTCTCCTTTACTCATggtaaaaaaacctcactaaaattatg; P7:GACGTCCAAACGCAAGTCCTGctgaaacctcaggcatttgag; P8:CATGGATGAGCTCTACAAATAAgcgcaacgcaattaatgtaag; P9:CATAATTTTAGTGAGGTTTTTTTACCatgagtaaaggagaagaacttttc; P10:CTTACATTAATTGCGTTGCGCttatttgtagagctcatccatg; P11:CATTAGCGTTATAGTGAATCCGCtaacgttctcagggcggggtg; P12:GAAAAGTTCTTCTCCTTTACTCATgcgacctcccgtttttccgcc; P13:CATTAGCGTTATAGTGAATCCGCtaaaacccatcgcttcagggc; P14:GAAAAGTTCTTCTCCTTTACTCATaatgaaacgctctcgtaagaatac; P15:CACCCCGCCCTGAGAACGTTAgcggattcactataacgctaatg; P16:GGCGGAAAAACGGGAGGTCGCatgagtaaaggagaagaacttttc; P17:GCCCTGAAGCGATGGGTTTTAgcggattcactataacgctaatg; P18:GTATTCTTACGAGAGCGTTTCATTatgagtaaaggagaagaacttttc; AbRFN-ribB_RED forward: ttgcatcagtcctgaaatgttcaaccgtattcttacgagagcgtttcattatgaatcagacgctactttc; PaRFN-ribB_RED forward: gtcgcgccggccatgctgcgcgcctgtgcggcggaaaaacgggaggtcgcatgaatcagacgctactttc, Ab/Pa RFN-ribB_RED reverse: agatcccggtgcctaatgagtgagctaacttacattaattgcgttgcgcgctggctttacgctcatgtg; yqiC_RED forward: caggacttgcgtttggacgtcgaactcttcacggcttacaaggtcgaggcgcgtcagctgcgcttgtagg; ribB_RED reverse: gattaaggcagtaaattaagcagcggttttcagctggctttacgctcatgtgcctgacggtatgccacca; yqiC seq. reverse: agttcgctgattctttgttc; ribB seq. reverse 1: agcggaattaacatcttgc; ribB seq. reverse 2: gcttcaatggtcacggtaa. Transition from capital to lowercase letters for P5–P18 denotes boundaries of fragments that facilitate in-fusion cloning. EcFMN–GFP reporter plasmids were constructed by fusing the EcFMN region, inclusive of 550 bp upstream of ribB through the start codon, to gfpuv and cloning into a vector with the low copy CloDF13 origin of replication. Primers P5 and P6 were used to amplify the EcFMN region from wild-type and resistant mutants by colony PCR, primers P7 and P8 were used to amplify the CloDF13 origin and SmR cassette from pCDF-1b (EMD Millipore), and primers P9 and P10 were used to amplify gfpuv from pGFPuv (Clontech). Upon purification, all three linear PCR products were combined using the in-fusion HD cloning system (Clontech) and transformed into TOP10 cells (Life Technologies) with selection on spectinomycin (MP biomedicals) to yield pCDF-EcFMN–GFP reporter plasmids. Plasmids were subsequently transformed into the MB5746 ribocilR mutant M5 background for compound testing. Initial attempts to create Pseudomonas aeruginosa and Acinetobacter baumannii FMN–GFP reporters in a similar fashion to the EcFMN–GFP reporters by using the PaFMN or AbFMN region, inclusive of 550 bp upstream of ParibE or AbribB through the start codon, did not yield constructs with sufficient baseline fluorescence (data not shown). In order to optimize fluorescence, hybrid constructs were made in which the E. coli promoter region was placed upstream of the PaFMN or AbFMN elements. Primer combinations P11/P12 or P13/P14 were used to amplify the PaFMN or AbFMN elements, respectively, from wild-type cells by colony PCR, and primer pairs P15/P16 and P17/P18 were used to amplify the E. coli promoter, gfpuv, and vector backbone from the previously constructed pCDF-EcFMN–GFP plasmid for combination with PaFMN and AbFMN, respectively. Purified linear PCR products were combined and transformed as described above to yield pCDF-EcPro-PaFMN–GFP and pCDF-EcPro-AbFMN–GFP reporter plasmids. Again, plasmids were transformed into the MB5746 ribocilR mutant M5 background for compound testing. The native, chromosomal E. coli ribB riboswitch was replaced with that of either A. baumannii or P. aeruginosa using a two-step λ-RED recombineering process44. In the first recombineering event, the GFPuv coding sequence from either the pCDF-EcPro-AbFMN–GFP or pCDF-EcPro-PaFMN–GFP plasmid was replaced with the E. coli ribB coding sequence (EcribB). To this end, MB5746 ribB::kan cells were grown in CAMH broth supplemented with 4 μg ml−1 riboflavin (reconstituted in 1:1 dH2O:ethanol) and transformed (as described below) with the temperature-sensitive plasmid pKD4644. Reactions were plated onto CAMH agar +50 μg ml−1 ampicillin at 30 °C. Next, either the pCDF-EcPro-AbFMN–GFP or pCDF-EcPro-PaFMN–GFP plasmid was transformed into MB5746 ribB::kan/pKD46 and plated onto CAMH + 4 μg ml−1 riboflavin + 50 μg ml−1 spectinomycin + 50 μg ml−1 ampicillin at 30 °C to maintain double plasmid selection. The resulting strains were recombineered with EcribB PCR product containing flanking regions homologous to the cognate pCDF plasmid. Substrate PCR products were obtained through colony PCR of the wild-type ribB locus of MB5746 and either the AbFMN-ribB_RED forward or PaFMN-ribB_RED forward primer in combination with the Ab/Pa FMN-ribB_RED reverse primer. Cells were recovered in CAMH broth for 1 h and Rib+ colonies were selected on CAMH agar +50 μg ml−1 spectinomycin and incubated at 37 °C to remove pKD46. The resulting strains, MB5746 ribB::kan (pCDF-EcPro-AbFMN-EcribB) or (pCDF-EcPro-PaFMN-EcribB) carry a plasmid-borne EcribB gene downstream of the native E. coli ribB promoter fused to either AbFMN or PaFMN. In the second recombineering event, the plasmid-borne AbFMN- or PaFMN-EcribB fusions engineered above were introduced into the E. coli chromosome in single-copy at the native ribB locus. MB5746 ribB::kan/pKD46 cells were grown to exponential phase in CAMH broth + 4 μg ml−1 riboflavin + 50 μg ml−1 ampicillin and electroporated with either AbFMN-EcribB or PaFMN-EcribB PCR product containing flanking regions homologous to the native ribB locus. These PCR products were amplified from pCDF-EcPro-AbFMN-EcribB or pCDF-EcPro-PaFMN-EcribB using the yqiC_RED forward and ribB_RED reverse primers. Unlike strains carrying the plasmid-borne hybrid EcribB constructs, the chromosomal hybrid fusion constructs do not yield enough riboflavin for optimal growth on CAMH in single copy. Therefore, reactions were recovered in CAMH broth containing very low levels of riboflavin (0.4 μg ml−1, a concentration that does not permit growth of the ribB deletion mutant), plated onto CAMH agar + 0.4 μg ml−1 riboflavin, and incubated at 37 °C to remove the pKD46 plasmid. The E. coli ribB promoter, hybrid riboswitch, and EcribB coding regions were sequenced in resulting Rib+ cells and additionally sequenced at joint sequences using yqiC seq. reverse, ribB seq. reverse 1, and ribB seq. reverse 2 primers. All transformations were electroporation reactions performed as suggested2 with some modifications. Around 30–50 ml of cells were grown in CAMH to exponential phase. For recombineering reactions, strains harbouring pKD46 were induced for 1 h with 1% arabinose before harvesting of cells. Cells were washed with 30 ml of ice-cold ddH O and pelleted at 4 °C, 3,000g for 10 min, followed by two additional washes with 1 ml ice-cold ddH O and pelleted each time at 8,000g at 4 °C for 2 min. Pellets were re-suspended in 300 μl ddH O and 100 μl cells were incubated with 1–2 μl PCR product for 5 min before electroporation. Electroporation reactions were performed using 0.1-cm gap cuvettes and a GenePulser II (BioRad) with settings at 200 Ω, 25 μF, and 1.8 kV. Cells were recovered at the appropriate temperature in 1 ml CAMH broth as described above and plated on CAMH agar containing the appropriate supplements. All PCR reactions were performed using GoTaq Green Master Mix (Promega Corporation) according to manufacturer’s instructions. DNA sequencing was performed by Genewiz, Inc. Crystals of the F. nucleatum FMN riboswitch in presence of the ligand were obtained following published protocols13, 45 with minor modifications. The RNA was synthesized in two strands: GGAUCUUCGGGGCAGGGUGAAAUUCCCGACCGGUGGUAUAGUCCACGAAAGCUU and GCUUUGAUUUGGUGAAAUUCCAAAACCGACAGUAGAGUCUGGAUGAGAGAAGAUUC. The oligonucleotides were purchased from Sigma-Aldrich. After reception each strand was dissolved in water, aliquoted so that each aliquot would contain the material necessary to make a 25 μl solution at 0.4 mM concentration. The aliquots were lyophylized using a Centrivap concentrator (Labconco) and kept at −20 °C for long-term storage. Prior to annealing the nucleic acids were re-suspended in 25 μl annealing buffer (10 mM cacodylic acid, 100 mM acetate, 4 mM MgCl adjusted to pH 6.8 using KOH). The oligomers were mixed together, along with 1.0 μl of an inhibitor stock solution at 50 mM in 100% deuterated DMSO, and annealed in a thermocycler by incubation at 37 °C for 30 min followed by cooling from 37 °C to 4 °C at a rate of 3 °C per min. The crystals were grown by vapour diffusion using a 15-well EasyXtal DropGuard X-Seal tool (Qiagen) after mixing 3 μl of riboswitch–ligand solution with 3 μl precipitant (0.1 M Na acetate, pH 5.0, 0.2 M MgCl , and 7 to 11% v/v PEG 4K). Small nuclei appear after a few days, and are made to grow larger for diffraction studies by controlled drying. Drying is achieved by substitution once a day of the adequate volume of well solution with a 50% v/v PEG 4K stock solution. The volume is calculated to achieve a ~2% increase in precipitant concentration per step. The crystals after growth are harvested and dipped for 1 to 2 min in a cryoprotectant solution (0.1 M Na MES, pH 6.5, 0.2 M MgCl , 10% v/v PEG 4K, 20% v/v glycerol, and ligand diluted to 1 mM concentration). Crystals were harvested with a mesh Litholoop (Molecular Dimensions Ltd) and flash-frozen in liquid nitrogen. X-ray diffraction data (Extended Data Table 2) were collected at the Advanced Photon Source (APS) sector 17 (IMCA) at 1.0 Å wavelength using a Pilatus 6M (Dectris) pixel array detector. 720 frames with an oscillation of 0.25° each were collected. The data were processed using the automated pipeline autoPROC46, with calls to the programs XDS47 for integration and AIMLESS48 for scaling. The structure was determined using PDB entry 3F4E as a starting point after removing all heterogeneous atoms including the FMN. The structure was refined without inclusion of the ligand coordinates at any step before and until an omit map difference map is generated to fit the compound. The steps include refinement using the program autoBUSTER49, corrections of the model and inclusions of several cations with Coot, and Cartesian simulated annealing using the program Phenix50 to further eliminate the potential of bias against FMN which was present in the PDB 3F4E entry. The set of ‘free’ reflections was taken from the same PDB entry 3F4E and completed as required. All refinement calculations after adding the ligand were performed using the program autoBUSTER49. Model visualization and rebuilding was performed using the program Coot51. All figures in the manuscript generated with PyMol52. Co-crystallization of the heptamer with ribocil was performed using a racemic mixture of the ligand. In spite of its limited resolution, 2.9 Å, the (R) isomer fits distinctly better than the (S) isomer in the initial electron density. Consistent with this observation, crystallographic refinement of a model which starts with the wrong (R) isomer ends up with a structure with the chiral centre inversed and nearly planar, an impossible stereochemistry. By contrast, the chiral volume remains unchanged in the course of refinement when starting from the (S) isomer. Notwithstanding the electron density map, due to the constrained nature of the binding site it is not possible to fit the (R) isomer while maintaining reasonable ligand stereochemistry and parallelism of the pyrimidynonyl and the methylaminopyrimidynyl between the bases of A48 and A85, and against the base of G62. Altogether, crystallographic and stereochemical considerations strongly support the conclusion that only the (S) form binds to the FMN aptamer. Further observations made later when the isomers were separated agree with this interpretation: only one of them is active against the riboswitch, and the ligand with the correct chirality is more active than the racemic mixture (Extended Data Fig. 3 and Extended Data Table 1). A homology model of the E. coli FMN aptamer was constructed using program mutate_bases53 of the 3DNA package using the F. nucleatum impX riboswitch aptamer X-ray structure as the template and the FMN aptamer alignment of E. coli, F. nucleatum, P. aeruginosa and A. baumannii (Extended Data Fig. 5). All nucleotide insertions in the E. coli sequence were removed in the model (Extended Data Fig. 5). There are 34 base changes among the 111 nucleotides modelled. Base pairing when present remains consistent. Energy minimization at A92 was performed to avoid VDW clashes using Macromodel (Schrodinger, LLC). Reporter strains were diluted to ~5 × 106 CFU ml−1 in CAMHB supplemented with 30 µg ml−1 spectinomycin. Compounds to be tested were serially diluted twofold through 11 points in 100% DMSO. A BioMek FX liquid handler was used to deliver 49 μl of diluted culture into a 384-well, black/clear-bottom assay plate followed by 1 μl of titrated compound. DMSO and antibiotic controls were added manually to appropriate wells and the plates were shaken for 1 min before incubating at 37 °C. After overnight growth, fluorescence, using 405 nm excitation and 510 nm emission, and absorbance at 600 nm, was assessed on an EnVision multiplate reader (Perkin Elmer). Fluorescence response (RFU), relative to full growth and fully inhibited (50 μM ribocil) controls and absorbance response (OD ), relative to full growth and sterile controls, were fitted to four parameter (variable slope) curves. The concentration of compound which decreased the specific fluorescence signal by 50% is reported as the GFP EC . Aptamers were first re-annealed at a 20 μM concentration in 4 mM KH PO , 16 mM K HPO , 64 mM KCl and 0.1 mM EDTA, pH 7.4 buffer by heating at 95 °C for 5 min followed by incubation at room temperature for 15 min. Only one re-annealing cycle was performed per aptamer sample. A 1.25-fold serial dilution of the re-annealed aptamer was prepared to have a final concentration ranging from 6.6 to 150 nM in 50 mM Tris-HCl, 100 mM KCl and 2 mM MgCl assay buffer, pH 7.4. This was mixed with FMN ligand (30–240 nM final concentration). Fluorescence signal was read using the Spectramax M5 at an excitation wavelength of 455 nm and emission wavelength at 525 nm with cut-off filter at 515 nm. The instrument was set up in kinetic mode to acquire data every 20 s. The steady-state K and the binding-competent fraction of aptamer were determined from fluorescence data obtained at 120 min of the reaction by fitting to a quadratic equation fully describing the binding equilibrium under tight-binding conditions. A twofold serial dilution of compounds was prepared to have a final assay concentration range from 1.22 to 10,000 nM. This was prepared in 50 mM Tris-HCl, 100 mM KCl and 2 mM MgCl assay buffer, pH 7.4, with 0.2% DMSO. FMN ligand concentration was 60 nM and the E. coli FMN aptamer concentration was 48 nM or, for ribocil, 150 nM. The fluorescence signal was read on the Spectramax M5 as described above. The steady-state binding competition data at 120 min was fitted to a cubic equation fully describing the competition binding equilibria to derive the K value for the compound, while fixing K to the value obtained earlier for FMN binding. The binding kinetics data was fitted by KinTek Explorer-based numerical integration with K constrained to derive the dissociation rate constant (k ). The association (k ) rate constant is then calculated from K and k . The Click-iT EdU Alexa Fluor 488 HCS assay kit (Life Technologies, C10351) was used to assess the potential cytoxicity of ribocil in mammalian cell cultures using a modified version of the manufacturer’s protocol. For the assay, mycoplasma-tested HeLa cells (ATTC) were seeded at 4,000 cells per well in 384-well poly-d-lysine-coated plates (Greiner, 781946) in 25 μl of culture medium (Optimem I, Life Technologies) and treated with a 20-point twofold dilution series of ribocil. After addition of EdU (5 μM) and incubation (37 °C) for 24 h, images were captured and analysed using an Acumen eXC3 (TTP Labtech Ltd) laser scanning cytometer. Total cell numbers were determined using Hoechst 33342 (Life Tech, H3530). Ribocil displayed no HeLa cell cytoxicity as detected by cell count (EC ≥ 100 μm, activity at 100 μM = 17%) or EdU measurement (EC ≥ 100 μM, activity at 100 μM = 11%).

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