Research ArticleTuberculosis

Structural Basis for Benzothiazinone-Mediated Killing of Mycobacterium tuberculosis

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Science Translational Medicine  05 Sep 2012:
Vol. 4, Issue 150, pp. 150ra121
DOI: 10.1126/scitranslmed.3004395


The benzothiazinone BTZ043 is a tuberculosis drug candidate with nanomolar whole-cell activity. BTZ043 targets the DprE1 catalytic component of the essential enzyme decaprenylphosphoryl-β-d-ribofuranose-2′-epimerase, thus blocking biosynthesis of arabinans, vital components of mycobacterial cell walls. Crystal structures of DprE1, in its native form and in a complex with BTZ043, reveal formation of a semimercaptal adduct between the drug and an active-site cysteine, as well as contacts to a neighboring catalytic lysine residue. Kinetic studies confirm that BTZ043 is a mechanism-based, covalent inhibitor. This explains the exquisite potency of BTZ043, which, when fluorescently labeled, localizes DprE1 at the poles of growing bacteria. Menaquinone can reoxidize the flavin adenine dinucleotide cofactor in DprE1 and may be the natural electron acceptor for this reaction in the mycobacterium. Our structural and kinetic analysis provides both insight into a critical epimerization reaction and a platform for structure-based design of improved inhibitors.


Tuberculosis (TB) represents a formidable challenge to global health and results from infection with the airborne pathogen Mycobacterium tuberculosis. In recent decades, synergy with the HIV/AIDS pandemic, widespread social instability, and poverty have all contributed to a resurgence of TB that increasingly manifests in multidrug-resistant (MDR) and extensively drug-resistant (XDR) forms (1, 2). Faced with the specter of untreatable disease, intensive efforts have been made to discover new drugs to replace the existing TB treatment that was developed in the 1960s. New TB drugs should be highly potent to reduce therapy duration and should inhibit new targets to ensure activity against both drug-susceptible and drug-resistant strains of M. tuberculosis in circulation.

A portfolio of promising new antitubercular agents is on the horizon (3, 4). Among these, the benzothiazinones (BTZs) represent a class of nitroaromatic molecules that kill M. tuberculosis cells in vitro, ex vivo, and in animal models of the disease (5). BTZ043, currently in the late stages of preclinical development (4), exhibits a minimal inhibitory concentration (MIC) of 1 ng/ml against M. tuberculosis, making it much more potent than any of the currently used drugs (Fig. 1A). The cellular target of BTZs is the DprE1 component of the mycobacterial decaprenylphosphoryl-β-d-ribofuranose-2′-epimerase encoded by the neighboring genes dprE1 and dprE2 (5), which are essential for growth of M. tuberculosis and Mycobacterium smegmatis (6, 7). Concerted expression of DprE1 and DprE2 is required to carry out the epimerization reaction that converts decaprenylphosphoryl-β-d-ribofuranose (DPR) into decaprenylphosphoryl-β-d-arabinose (DPA) (Fig. 1B) (8). DPA is the sole precursor for the synthesis of the arabinan moiety of the mycobacterial cell wall, whose unique composition renders mycobacteria insensitive to a number of antibiotics (9). DprE1 from most actinobacteria is susceptible to BTZ043 (5). Furthermore, all M. tuberculosis clinical isolates tested so far, from drug-sensitive, MDR, and XDR TB cases, are susceptible to BTZ043, making this drug a promising candidate for the treatment of all forms of TB and possibly for other mycobacterial diseases, such as leprosy (10). More recently, two other families of less potent antitubercular compounds, namely, the dinitrobenzamides such as DNB1 (MIC, 0.072 μg/ml) and benzoquinoxalines such as VI-9376 (MIC, 1 μg/ml) (Fig. 1A), were also found to target DPR epimerization, and several other scaffolds targeting this function are being developed (11, 12).

Fig. 1

Inhibitors and enzymatic activity of DprE1. (A) Structures of antitubercular compound families that target DprE1. BTZ043 (MIC, 1 ng/ml) is in late preclinical development (5). Reduced BTZ043 analogs BTZ045 (amino) and BTZ046 (hydroxylamino) show MIC values >500-fold higher than that of BTZ043. DNB1 represents the dinitrobenzamide family of inhibitors (11) (MIC, 0.072 μg/ml or 0.02 μM). VI-9376, a benzoquinoxaline, was also reported to target DprE1 (12) (MIC, 1 μg/ml or 2.9 μM). (B) Epimerization reaction on the 2′-hydroxyl group of DPR, catalyzed by the mycobacterial DprE1/DprE2. DPR is converted into DPA, an essential precursor for the synthesis of the arabinan moiety of the mycobacterial cell wall (9). DprE1 catalyzes the first step through a FAD-dependent process that requires an electron acceptor for enzyme turnover, which in vitro can be either molecular oxygen, DCPIP, or menaquinone (MQ). BTZ043 is converted to a nitroso derivative by DprE1-reduced flavin cofactor (13).

The epimerization of DPR to DPA takes place in two sequential oxidation-reduction reactions. First, DprE1 oxidizes DPR to decaprenylphosphoryl-2-keto-β-d-erythro-pentofuranose (DPX), which is then reduced by DprE2 to DPA (Fig. 1B) (13). Genetic analyses of resistant mutants restricted the molecular target of BTZs (as well as that of DNB1 and VI-9376; Fig. 1A) to the DprE1 component of the epimerase (5, 14). In particular, point mutations at a cysteine residue (Cys387 in M. tuberculosis, Cys394 in M. smegmatis) drastically increased the MIC of BTZ043, by up to 10,000-fold, highlighting the potential of DprE1 as a truly selective drug target (15). Mass spectrometric analysis of M. tuberculosis DprE1 overexpressed in M. smegmatis cells treated with BTZ043 demonstrated that the inhibitor is a prodrug, which is activated inside the cell to a nitroso derivative that covalently reacts with a cysteine residue on the target protein (16). The covalent nature of BTZ043 inhibition was further confirmed in vitro with purified recombinant M. smegmatis DprE1 (13).

DprE1, a 51-kD protein highly conserved among mycobacteria, shows 83% sequence identity between the M. tuberculosis and M. smegmatis orthologs (fig. S1). DprE1 shares moderate sequence similarity to flavoenzymes of the vanillyl-alcohol oxidase class (17), and on this basis, it was predicted to act as a decaprenylphosphoryl-β-d-ribofuranose-2′-oxidoreductase through a flavin adenine dinucleotide (FAD)–dependent mechanism. Here, we present the crystal structure of M. smegmatis DprE1 in its native form and in a complex with the BTZ043 inhibitor, revealing the mechanism for covalent inhibition. In addition, we report the biochemical analysis of both wild-type and mutant forms of the protein, and use a fluorescent BTZ analog to probe the subcellular localization of DprE1.


DprE1 structure reveals a flavoenzyme two-domain topology

Numerous attempts to produce M. tuberculosis DprE1 were made using multiple constructs and expression systems, but these yielded insoluble or inactive enzyme (see Supplementary Methods and fig. S2). Consequently, we focused on M. smegmatis DprE1 that was produced in soluble form with good yields from a pET SUMO construct (His6-SUMO tag) or from a pET32b construct (thioredoxin-His6 tag). Cleavage of the protein tags by specific proteases and subsequent purification afforded DprE1 of high purity (fig. S3), which was used for crystallization trials and activity assays.

The crystal structure of native M. smegmatis DprE1 in complex with its FAD cofactor was solved at 2.1 Å-resolution (Table 1 and Fig. 2A). Molecular replacement and energy-optimized rebuilding procedures by the Phaser and Rosetta programs were used to obtain an initial DprE1 model starting from the coordinates of cytokinin dehydrogenase [Protein Data Bank (PDB) code 1w1o] (18). Despite the low homology of the starting model (17% sequence identity), the high-quality x-ray data (Table 1) combined with powerful computational algorithms produced an excellent electron density map (Fig. 2B). This yielded an almost complete initial DprE1 model, which, upon further manual building and refinement, gave the final structure of DprE1. The structure obtained from the 468–amino acid DprE1 protein does not include the first 13 residues and two other segments (residues 275 to 303 and 330 to 336), which are not visible in the electron density map, probably because they either are disordered or adopt multiple conformations (Fig. 2A). SDS–polyacrylamide gel electrophoresis (SDS-PAGE) analysis on dissolved crystals using silver staining ruled out possible proteolytic events.

Table 1

Data collection and refinement statistics for DprE1 structures. One crystal was used for each structure. Values in parentheses are for the highest-resolution shell.

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Fig. 2

Crystal structure of M. smegmatis DprE1. (A) Light blue ribbon diagram of the DprE1 overall structure with labeled N and C termini. Disordered regions are indicated by dashed lines (numbers in parentheses correspond to the residues that flank the disordered parts). The FAD cofactor is represented in yellow, with nitrogen, oxygen, and phosphorus atoms colored in blue, red, and magenta, respectively. The active-site cavity is shown as a pink surface. (B) Refined 2mFo-DFc electron density map (contoured at 1.2σ) for the FAD cofactor and the Cys394 known to be the target of covalent modification by BTZ043. For clarity, electron density contours farther than 3 Å from atoms in the figure have been omitted. Orientation and color code are as in (A), except for protein carbon atoms, which are in light blue; sulfur atom is in green. (C) Close-up view of the DprE1 active site. Molecule orientation and color code are as in (B). Water molecules are drawn as red spheres. Black dashed lines indicate H bonds. The loop formed by residues 316 to 329 is shown, which shields the active-site cavity. (D) Electrostatic surface potential of DprE1 showing positively and negatively charged areas in blue and red, respectively. The FAD cofactor is visible in semitransparency [color code as in (B)]. Loop 316 to 329 is also visible in semitransparency.

The DprE1 structure features the common two-domain topology (Fig. 2A) observed in other flavoenzymes such as alditol oxidase and cytokinin dehydrogenase [root mean square deviation (RMSD) values calculated for the 345 Cα atoms of the aligned residues are 2.1 and 2.5 Å, respectively] (18, 19). The isoalloxazine ring of the noncovalently bound FAD lies at the interface between the cofactor-binding domain and the substrate-binding domain, where the two disordered regions are also located. The prosthetic group borders the active-site cavity of about 130 Å3, which includes the critical Cys394 (Cys387 in M. tuberculosis DprE1; Fig. 2, B and C) (13). Despite the structural and functional divergence, the DprE1 active site shares similarity with that of alditol oxidase (19). In particular, His139, Gly140, Gln341, and Lys425 (Fig. 2C) are conserved, whereas Tyr67, Gln343, and Lys374 are replaced by relatively conserved residues (Phe, Glu, and Arg, respectively). This similarity reflects the common chemical activity of the two enzymes because both oxidize a C-OH group in a sugar substrate. The active-site cavity is shielded by a loop formed by residues 316 to 329, which reside on the protein surface and are followed by one of the two disordered regions of the structure (Fig. 2, A and C). The protein surface located close to this loop and belonging to the FAD-binding domain is sprinkled with a cluster of positively charged residues (Fig. 2D), which might be involved in interactions with the cell membrane.

To assess the main differences between the DprE1 proteins of M. smegmatis and M. tuberculosis, we generated a homology model of the latter based on the M. smegmatis DprE1 structure using the SWISS-MODEL interface (2022). Given the high sequence identity between the two DprE1 proteins (83%), the model shows conservation of all residues in the active site, with more significant differences observed on the protein surface.

The DprE1 complex with BTZ043 shows a covalent adduct with Cys394

BTZ043 is a mechanism-based covalent inhibitor, which requires the enzymatic activity of the protein to reduce BTZ043 to the cysteine-reacting nitroso analog. Consequently, the DprE1-BTZ043 adduct had to be generated before crystallization trials. This was achieved by incubating DprE1 with BTZ043 and FPR (farnesylphosphoryl-β-d-ribofuranose), an analog of DPR with a shorter polyprenyl chain, which was shown to be a good enzyme substrate (13). Adduct formation was confirmed by mass spectrometry (fig. S4). Crystals of the DprE1-BTZ043 complex were obtained in conditions different from those that yielded crystals of the native protein. Inclusion of the ionic liquid tetrabutylphosphonium bromide in the crystallization condition proved crucial to generate good quality crystals of the BTZ043-labeled protein that diffracted to 2.6 Å (Table 1).

The overall structure of the DprE1-BTZ043 adduct is very similar to that of the native protein (RMSD of 0.285 Å for 346 Cα carbons), with the FAD cofactor bound exactly in the same conformation. The electron density map was of very good quality and unambiguously showed the presence of a covalent adduct between BTZ043 and Cys394 (Fig. 3, A and B). The covalent adduct causes Cys394 to adopt a rotamer conformation different from that observed in the native structure. BTZ043 binds in front of the FAD isoalloxazine ring with an angle of about 45° between the ring systems of the two molecules (Fig. 3A). The loop (residues 324 to 329) blocking the active-site cavity in the native structure (Fig. 2C) is partly disordered in the BTZ043-bound protein (Fig. 3C). Superposition to the native protein structure shows that movement of the loop is required to accommodate the BTZ043 molecule (Fig. 3B) and, most likely, also to bind the DPR substrate.

Fig. 3

Structure of DprE1 in complex with BTZ043. (A) Refined 2mFo-DFc electron density map provides clear evidence of the formation of the semimercaptal adduct between the reduced BTZ043 form and Cys394 of DprE1. For clarity, electron density contours farther than 3 Å from atoms in the figure have been omitted. The protein molecule is oriented as in the native structure in Fig. 2, A and B. Nitrogen, oxygen, sulfur, phosphorus, and fluoride atoms are colored in blue, red, green, magenta, and cyan, respectively. Carbon atoms are represented in yellow in the FAD cofactor, black in BTZ043, and gray in the protein. (B) Surface representation showing the DprE1 active site with BTZ043 bound to Cys394 in front of the FAD cofactor. Color code is as in (A). The structure is rotated about 90° around an axis perpendicular to the plane of the figure with respect to the orientation in (A). Superposition with the native structure shows that a portion of the flexible loop shielding the active-site cavity (drawn in light blue as in Fig. 2C) is displaced by the BTZ043 molecule. (C) BTZ043 binding to DprE1 and interactions with active-site residues. Structure orientation is the same as in Fig. 2C. Color code is as in (A). Water molecules are represented as red spheres. Dashed lines represent H bonds. The visible portion (residues 316 to 323) of the cavity loop is shown in gray.

Comparison with the native structure (Fig. 2C) shows that two water molecules occupying the inner part of the cavity are displaced to a different position in front of the flavin ring, with one of them bridging a network of hydrogen bonds between the inhibitor and Tyr67. Moreover, a key interaction is observed between the semimercaptal hydroxyl group and the side chain of Lys425 (Fig. 3C). The CF3 group of BTZ043 is well placed in a small pocket lined by His139, Gly140, Lys141, Lys374, and Phe376 and interacts with the amide of Asn392 (Fig. 3C). Notably, no major interactions are observed with other parts of BTZ043, except for a weak hydrophobic interaction between the side chain of Leu370 and the piperidine ring of BTZ043. The spirocyclic moiety of BTZ043 is located at the protein surface and lacks full electron density, namely, to account for its methyl group (Fig. 3A). As indicated by the mass spectrometry data, the protein used for crystallization was intact and contained the full BTZ043 semimercaptal adduct (fig. S4), suggesting that the partially undefined electron density is due to disorder and/or multiple conformations, rather than to degradation.

DprE1: An oxidase or a dehydrogenase?

The oxidative reaction catalyzed by DprE1 at the 2′-hydroxyl group of the DPR substrate implies that its FAD cofactor undergoes a reductive half-reaction generating reduced flavin (FADH2), which has to be reoxidized by an electron acceptor to start a new catalytic cycle (Fig. 1B). We verified that DprE1 can use molecular oxygen as electron acceptor, using a horseradish peroxidase (HRP)–coupled Amplex Red assay that probes for hydrogen peroxide production due to flavin reoxidation by oxygen (23) in the presence of the substrate FPR. We also found that DprE1 activity increases by about 30% when excess FAD is present, suggesting that a major fraction of the protein molecules is in the apo form. This fact is in agreement with our observation that DprE1 tends to lose FAD during purification. Initial kinetic data were obtained at 25°C, and the initial velocity versus substrate concentration plot resulted in a sigmoidal curve that was best fitted with the Hill equation (Table 2). This was unexpected because DprE1 is not predicted to be an allosteric enzyme, and gel filtration chromatography indicated that the protein is monomeric, ruling out activation mechanisms due to oligomerization. The sigmoidal shape of the steady-state kinetics is less pronounced when experiments are performed at 37°C with essentially unchanged kinetic constants.

Table 2

Steady-state kinetics of M. smegmatis DprE1 enzymes.

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We then probed other known electron acceptors to assess whether these could replace molecular oxygen in reoxidizing the reduced FAD in DprE1. We observed that 2,6-dichlorophenolindophenol (DCPIP) (24) is an efficient electron acceptor substrate for DprE1, showing a fourfold increase in the value of kcat compared with that measured for oxygen (Table 2). Furthermore, with DCPIP, the steady-state data could be fitted to the Michaelis-Menten equation without any indication of sigmoidal behavior. These findings indicate that DprE1 is unlikely to be a true oxidase but rather a dehydrogenase that reacts poorly with oxygen and likely uses an alternative electron acceptor in the bacterium such as menaquinone. We then tested whether this electron acceptor, present in mycobacterial membranes, could reoxidize the DprE1-bound FADH2 after its reduction by exposure to light and deazaflavin. On monitoring the ultraviolet-visible (UV-Vis) absorbance spectrum, we observed full reduction of FAD to FADH2, which was promptly converted to its reoxidized form upon addition of menaquinone (Fig. 4A). Therefore, menaquinone could indeed be the physiological substrate for regenerating the oxidized form of DprE1 after each catalytic cycle.

Fig. 4

DprE1 inhibition by BTZs, DNB1, and VI-9376. (A) UV-Vis absorbance spectrum showing the anaerobic DprE1 flavin reoxidation carried out by menaquinone. AU, absorbance units. (B) UV-Vis absorbance spectrum showing the anaerobic DprE1 flavin reoxidation carried out by BTZ043. (C) Steady-state progress curves obtained for the inactivation of DprE1 by BTZ043 at different concentrations (reported on the right of each curve) with the HRP-coupled Amplex Red assay. The standard reaction mixture contained 20 mM glycylglycine (pH 8.5), 150 μM FPR, 0.3 μM DprE1, 50 μM Amplex Red, and 0.35 μM HRP, and the assay was performed at 37°C. Progress curves were fit to the single exponential equation for slow-binding inhibitors: product = v0[1 − exp(−kobst)]/kobs, where v0 is the initial velocity for each curve. The kobs values were then analyzed by the method of Kitz and Wilson to yield kinact and Ki(inact) (see inset) (41). (D) DprE1 initial velocities measured by the DCPIP assay plotted versus inhibitor concentration. In each measurement, the protein sample (3 μM) was incubated with 100 μM FPR and variable concentrations of the inhibitor (0 to 50 or 0 to 200 μM depending on the inhibitor). The IC50 values were determined by fitting the data as described in the Supplementary Methods. (E) Mechanism of DprE1 inhibition by BTZ043.

Site-directed mutagenesis identifies Lys425 as a key residue for DprE1 activity

After analysis of the active-site architecture and the DprE1 residues that interact with BTZ043, we generated three active-site mutants, namely, Cys394Gly, Gln343Ala, and Lys425Ala (Table 2), and characterized them by activity studies using the DCPIP functional assay. First, replacement of Cys394 with Gly had an evident effect on activity by causing a 14-fold decrease in the catalytic efficiency (kcat/Km). Similarly to the Cys394Gly variant, the Gln343Ala mutant is also enzymatically active, but displayed a marked decrease in catalytic efficiency (10-fold), indicating that this residue also plays a role in the active-site architecture and enzyme catalysis (Table 2).

Lys425 is homologous to a Lys residue (Lys375) present in the active site of alditol oxidase that is directly involved in catalysis, where it was proposed to activate the sugar substrate by interacting with the hydroxyl group oxidized by the enzyme (19). Strikingly, in DprE1, the Lys425Ala mutant was completely inactive (FAD cofactor was normally incorporated as indicated by its spectral properties), suggesting that Lys425 plays the same role in substrate binding and catalysis as in alditol oxidase. This is consistent with its position in front of the flavin (Fig. 2C), and therefore, this residue can be predicted to interact with the 2′-hydroxyl group of the substrate, the site of oxidation of DPR by DprE1 (Fig. 1B).

Nitroso-BTZ043 is generated upon reoxidation of DprE1-bound FADH2

With the DCPIP assay described above, we were able to perform an in-depth analysis of DprE1 inhibition by BTZ043. To verify that DprE1 is responsible for the activation of BTZ043 through its reduction to the nitroso analog (13, 16), we probed the ability of BTZ043 to reoxidize protein-bound FADH2 as described above for menaquinone. We verified that after full reduction of FAD to FADH2 as described, this was rapidly reoxidized upon addition of BTZ043 (Fig. 4B). Enzyme reoxidation was not observed with the analogs of BTZ043 (BTZ045 and BTZ046) lacking a nitro group. This is consistent with previous results (13), unequivocally demonstrating that BTZ043 is a suicide inhibitor, activated in mycobacteria by reacting with reduced DprE1 and then forming the semimercaptal covalent adduct with Cys394, as illustrated in Fig. 4E. We also performed a kinetic study of DprE1 inactivation by analysis of progress curves at different BTZ043 concentrations (Fig. 4C). We used oxygen as electron-accepting substrate instead of DCPIP because the latter is known to interact with Cys residues over a certain time frame (25) and could interfere with the formation of the BTZ043-Cys394 adduct. The assays were performed at 37°C to reduce the sigmoidal behavior of the progress curves, which could then be fitted to the single exponential equation (Fig. 4C). The Ki(inact) and kinact for the inhibition of DprE1 by BTZ043 were measured to be 12.5 μM and 0.58 min−1, indicating that covalent inactivation of DprE1 by BTZ043 occurs very slowly.

Remarkably, the noncovalent BTZ045 and BTZ046 inhibitors exhibit IC50 (half-maximal inhibitory concentration) values that are not drastically different from that of BTZ043: 4.5 μM for BTZ043, 11.0 μM for BTZ045, and 19.7 μM for BTZ046 (measured by the DCPIP assay with a 7-min incubation time for the enzyme-inhibitor complex; Fig. 4D). Nevertheless, their different mechanisms of inhibition translate into widely different antibacterial activities as discussed below.

Dinitrobenzamides and nitrobenzoquinoxalines are also suicide inhibitors

The dinitrobenzamides and the nitrobenzoquinoxaline VI-9376, initially discovered in whole-cell screens for antitubercular activity, also target DprE1 (11, 12). To investigate the mechanism of inhibition by these compounds, we attempted co-crystallization with DprE1 using the procedure described above for BTZ043. However, despite considerable efforts, high diffraction grade crystals have not yet been obtained. Therefore, we took advantage of the DprE1 crystal structure and performed docking studies with DNB1 and VI-9376 to generate models of the complexes formed between DprE1 and these compounds. The docking results suggest that both compounds indeed fit nicely in the BTZ043-binding site and make the key interactions observed for BTZ043 (fig. S5). The weaker inhibitory activity of VI-9376 might be due to its small size and the limited number of functional groups, therefore exhibiting only weak interactions with the active site. DNB1, being a more flexible molecule, was seen to adopt a bent conformation in the docked pose, exploring space available in the active site that was not occupied by BTZ043.

We further characterized the DprE1 inhibition by these compounds through enzyme activity studies and mass spectrometry analysis. Mass spectrometry confirmed that DNB1 (dinitrobenzamide selected for this study) and VI-9376 also form a covalent adduct with DprE1 (fig. S4). Enzyme labeling reactions were performed in parallel and in the same conditions for BTZ043, DNB1, and VI-9376. Whereas for BTZ043 full labeling of DprE1 was observed, for DNB1 and VI-9376 only partial labeling was achieved (67 and 53%, respectively). Furthermore, the masses observed for the DNB1- and VI-9376–DprE1 adducts were different from those expected based solely on the formation of the semimercaptal product, indicating secondary structural modifications. Finally, the IC50 values determined for DNB1 (15.8 μM) and VI-9376 (57.4 μM) were 3.5- and 12.8-fold higher, respectively, than that determined for BTZ043 (Fig. 4D). Collectively, these data show that DNB1 and VI-9376 are also suicide inhibitors of DprE1 and reflect the weaker antimycobacterial activities of these two compounds compared with that of BTZ043.

Fluorescently labeled BTZ localizes DprE1 in the poles of live M. tuberculosis cells

Given that BTZ043 covalently binds to recombinant DprE1 very efficiently, we wondered whether a labeled BTZ could be used in live mycobacteria to probe the subcellular localization of this protein and consequently gain further insight into the cidality of this family of compounds. This could provide valuable information regarding uptake of DprE1 inhibitors by mycobacteria and indicate whether the protein is localized in the cytosol or the cell membranes.

We synthesized a fluorescent BTZ043 analog labeled with TAMRA (BTZ-TAMRA; Fig. 5 and fig. S6; synthesis described in the Supplementary Materials). To avoid clashes of the fluorophore with the active site and allow efficient binding, we connected TAMRA directly to the BTZ ring through a flexible tri-PEG (polyethylene glycol) linker. This compound was tested for antimycobacterial activity and it still exhibited appreciable efficacy (6 μg/ml), taking into account the substantial structural differences with BTZ043, namely, the presence of the bulky TAMRA fluorophore. M. tuberculosis was incubated with BTZ-TAMRA at three different concentrations around the MIC value, and images were taken at different time points by fluorescence microscopy. The kinetic analysis showed that BTZ-TAMRA can enter the cells quickly and gives rise to highly fluorescent spots localized at the poles of the bacteria, thereby defining the subcellular localization of DprE1 (Fig. 5).

Fig. 5

Subcellular localization of DprE1 in M. tuberculosis, probed by BTZ-TAMRA, a fluorescently labeled BTZ analog. Exponential M. tuberculosis cultures were exposed to BTZ-TAMRA and observed by fluorescence microscopy at different time points. Images were obtained with excitation and emission wavelengths of 542 and 585 nm, respectively. Higher fluorescence intensity is observed at the poles of the cells. Scale bars, 2 μm.


Isoniazid (INH) and ethambutol, two of the four first-line TB drugs, inhibit cell wall biosynthesis, which remains an attractive target for drug development (26). However, replacement of ethambutol, a weak drug inhibiting the last step in the DPA biosynthetic pathway, by a more potent compound has been envisioned, and a possible candidate is BTZ043 (Fig. 1A), which is now in late-stage preclinical development. BTZ043 acts two steps upstream of ethambutol by preventing formation of DPA (Fig. 1B), the sole donor of arabinose for biosynthesis of the essential cell wall arabinan components. DprE1 has been widely acclaimed as a new tractable target for TB drug development and is one of the few to have been pharmacologically validated. Here, through structural and functional characterization of DprE1 and its inhibition by BTZ043 and other nitro-compounds, we present an advance in understanding the mechanism of action at the atomic level.

The major difference between the native and BTZ043-bound DprE1 structures resides in a loop segment (residues 323 to 329) that restricts access to the active site in the native protein. Superposition of the two structures shows that binding of BTZ043 and most likely of the DPR substrate to the active site requires a conformational change of this loop, which might then serve as a switch between active and inactive forms of the enzyme. DprE1 likely forms a membrane-associated complex with DprE2 (8); thus, the conformational change might take place upon formation of the complex, leading to DprE1 activation. Another disordered loop (residues 275 to 303) and the disordered N terminus of the protein are also candidates for protein-protein and protein-membrane interactions. The observed cluster of positively charged residues at the surface of the protein could provide additional interactions between DprE1 and the membrane surface. Further support for this hypothesis comes from the fact that DPR, the natural substrate of DprE1, contains a long hydrophobic decaprenyl chain and is embedded in the cell membrane. DprE1 is likely a dehydrogenase, and not an oxidase, because it uses organic electron acceptors more efficiently than molecular oxygen to regenerate its FAD cofactor. In particular, our biochemical data indicate that the membrane-embedded menaquinone may represent the natural electron acceptor for reoxidation of the FAD cofactor of DprE1 in mycobacteria.

Subcellular localization studies with the fluorescent BTZ-TAMRA provide a clear indication that DprE1 is located at the poles of mycobacterial cells, demonstrating that this is the site where arabinan biosynthesis occurs, as was previously reported for peptidoglycan and mycolic acid (27). Synthesis of all three major cell wall components is thus spatially coordinated.

DprE1 oxidative activity and the concomitant FAD reduction are essential for activation of BTZ043 and effective inhibition of the enzyme through the formation of a covalent adduct. Residues that make key interactions with BTZ043, in particular Cys394 and Lys425, are also critical for full enzyme activity. Notably, the mutation of Lys425 led to complete loss of DprE1 activity, showing that this residue plays a key role in the oxidation of the 2′-hydroxyl group of DPR. On the other hand, despite losing significant activity, the Cys394Gly enzyme remained functional, indicating that this residue is not strictly needed for catalysis. This is fully consistent with the viability of the naturally BTZ043-resistant mycobacteria M. aurum and M. avium, where Cys394 has been replaced by Ser or Ala in DprE1, respectively, and with the spontaneous resistant mutants of M. tuberculosis, M. smegmatis, or Mycobacterium bovis Bacille Calmette-Guérin (BCG) bearing Gly or Ser in this position (5). Decreased DprE1 activity may account for a slight defect in growth rate observed for the Cys mutants in M. tuberculosis, M. bovis BCG, and M. smegmatis (S.T.C., unpublished results).

The inactivation of DprE1 by BTZ043 is time-dependent, whereas inhibition by its reduced amino and hydroxylamino analogs (BTZ045 and BTZ046, respectively) is not. However, the respective IC50 values are similar, ranging from 4.5 to 20 μM, indicating that formation of the initial enzyme-inhibitor complex is likely to be the rate-limiting step in the inhibition process. Hence, the key difference between BTZ043 and its reduced forms lies in the steps that follow binding to the reduced enzyme: Whereas BTZ043 can be reduced to the nitroso analog and immediately react with the adjacent Cys394, thereby irreversibly inactivating the enzyme, BTZ045 and BTZ046 cannot undergo this reaction. This explains the marked difference observed in their antitubercular activity (5) and the loss of potency of BTZ043 against the Cys394 mutants: Being unable to form a covalent adduct with Cys394, BTZ043 becomes a weak inhibitor of DprE1, just like BTZ045 and BTZ046 with the wild-type enzyme. These inhibition studies also highlight a potential limitation of target-based screens using DprE1 because inhibitors like BTZ046 will be found that may not prove effective in killing mycobacteria.

The dinitrobenzamides and the nitrobenzoquinoxalines, two other families of compounds known to target DprE1 (11, 12), are also mechanism-based irreversible inhibitors. The observed slower labeling of DprE1 by DNB1 and VI-9376 detected by mass spectrometry indicates either that these are reduced less efficiently to their nitroso analog or that, once generated, this intermediate does not react with Cys394 as promptly as the nitroso-BTZ043. These observations and the higher IC50 values measured for DNB1 and VI-9376 agree with their weaker antimycobacterial activity when compared with BTZ043.

The structural and enzymatic characterization of DprE1 is crucial for deepening our understanding of this target and the mode of action of BTZ043. This is an important development for a drug that is on track to enter clinical studies (4) and for which no resistant M. tuberculosis clinical isolates have been found so far, even among MDR and XDR TB cases (10). These structural data may underpin the development of other drugs, for instance, through structure-based design, and enable rational exploration of the active site of the enzyme. Alternative inhibitor scaffolds for DprE1 could then be found, including noncovalent inhibitors, which might be less susceptible to the development of resistance due to mutation of the active-site cysteine targeted by BTZ. The potential of this approach is highlighted by the docking studies presented here, in particular for DNB1 that partly occupies a pocket adjacent to the BTZ043-binding site.

To our knowledge, this is only the second structure of a TB drug in complex with its target, after that of the INH-NADH (reduced form of nicotinamide adenine dinucleotide) adduct in the enoyl-(acyl-carrier-protein) reductase InhA (28). This is paradoxical given the importance of TB as a global health problem, and reflects in part both the dearth of good targets and the difficult nature of mycobacterial drug discovery.

Materials and Methods

Protein expression and purification

All chemicals were purchased from Sigma-Aldrich, unless otherwise stated. M. smegmatis dprE1 (mc2155 strain, msmeg_6382 gene) was cloned into a modified version of pET32b vector (Novagen) containing a PreScission protease cleavage site between the tag and DprE1 (13). All point mutants were generated with the QuikChange (Stratagene) polymerase chain reaction–based method on the pET32b construct. Mutant enzymes were purified with yields comparable to the wild type. The proteins expressed from the pET32b construct were produced and purified with a modified protocol (Supplementary Methods). Improvement in protein yield was obtained by recloning the gene in the pET SUMO vector (Invitrogen) both in the full-length form of the DprE1 protein and as an N-terminal truncated construct lacking the first five amino acids (DprE1-Δ6, fig. S1), which were predicted to be disordered by bioinformatics analysis. Recombinant proteins were expressed fused to an N-terminal His6-SUMO tag in Escherichia coli BL21-DE3 in LB medium at 25°C for 5 hours (after induction with 1 mM isopropyl-β-d-1-thiogalactopyranoside). Cells were harvested by centrifugation and resuspended in 50 mM sodium phosphate (pH 8.0), 300 mM NaCl, 5% (v/v) glycerol, and 1 mM phenylmethylsulfonyl fluoride [20 g of cells/100 ml of buffer with two complete EDTA-free protease inhibitor cocktail tablets (Roche)]. Cell disruption was carried out by EmulsiFlex-C3 (Avestin), and the extract was obtained by centrifuging at 70,000g for 45 min at 4°C. Protein purification was carried out on an ÄKTA Purifier fast protein liquid chromatography system. The cell extract was loaded onto a HisTrap (1 ml, GE Healthcare), and the protein was eluted with 100 mM imidazole. The eluted fractions were yellow, indicating that the purified protein incorporated the FAD cofactor. Tag cleavage was achieved by overnight incubation with 0.3 mg of SUMO protease dialyzed against resuspension buffer, followed by a second HisTrap purification step to remove both tag and SUMO protease. After this step, the protein tends to lose the FAD cofactor, and the sample was therefore incubated with 100 μM FAD for 1 hour before the following purification steps, namely, ion exchange chromatography with a MonoQ column (GE Healthcare) followed by gel filtration (Superdex200 HR 10/300, GE Healthcare) in 25 mM potassium phosphate buffer (pH 7.2) and 5% glycerol. Sample purity was checked by SDS-PAGE, and protein concentration was evaluated by UV-Vis absorbance spectrum. The extinction coefficient at 449 nm (related to the FAD cofactor) was experimentally determined to be 10,300 M−1 cm−1 for the full-length protein and 13,530 M−1 cm−1 for the DprE1-Δ6 construct. The Abs280/Abs449 ratio was used to evaluate the fraction of holoenzyme (that is, containing FAD) with respect to total protein.

Activity and inhibition studies

The enzyme activity of DprE1 was spectrophotometrically determined with FPR as substrate and DCPIP as electron acceptor. FPR was synthesized as previously reported (29). Enzyme assays were performed on a Varian Cary 100 UV-Vis spectrophotometer equipped with temperature-controlled cell holder (T = 25°C), measuring the decrease in absorbance of DCPIP at 600 nm (ε = 19,100 M−1 cm−1). The standard reaction mixture contained 20 mM glycylglycine buffer (pH 8.5), 500 μM FPR, and 50 μM DCPIP, and the reaction was started by adding the enzyme solution (final concentration, 0.3 μM). Alternatively, the reactivity with molecular oxygen was determined measuring the H2O2 formation by the HRP-coupled Amplex Red assay, following the formation of resorufin at 560 nm (ε = 54,000 M−1 cm−1) at 25°C (or 37°C for the progress curve analysis). The standard reaction mixture contained 20 mM glycylglycine (pH 8.5), 500 μM FPR, 50 μM Amplex Red, and 0.35 μM HRP. Steady-state kinetic studies were performed by assaying the activity at different FPR concentrations (10 to 500 μM). All measurements were performed in triplicate. The kinetic constants, Km and kcat, were determined fitting the data to the Hill equation with Origin 8 software.

For inhibition studies, DprE1 (3 μM) was incubated for 7 min with 100 μM FPR in the presence of different inhibitor concentrations (0 to 100 μM for BTZs and DNB1 inhibitors; 0 to 200 μM for VI-9376; all inhibitors were dissolved in 100% dimethyl sulfoxide whose concentration was kept constant in the incubation mixture). Then, the enzyme activity was measured with the DCPIP assay with a final protein concentration of 0.3 μM (see above). The IC50 values were obtained by plotting the initial velocities with the following equation:A[I]=A[0]×(1[I][I]+IC50)where A[I] is the enzyme activity at inhibitor concentration [I] and A[0] is the enzyme activity without inhibitor.

The ability of menaquinone and BTZs to reoxidize the flavin of DprE1 was determined as follows: DprE1 solution (10 μM) was made anaerobic in a rubber septum–sealed cuvette (Hellma) by alternating cycles of vacuum with flushing of O2-free argon. The enzyme was then photoreduced by light irradiation in the presence of 0.1 μM 5-deazaflavin, and the photoreaction was followed spectrophotometrically. BTZ043 or menaquinone (10 μM) was then added, and the flavin reoxidation was monitored spectrophotometrically.

Crystallization and structure determination

Two crystal forms of DprE1 were obtained with the recombinant protein produced from the pET32b and pET SUMO constructs. Crystals of the native protein were obtained with the DprE1-Δ6 pET SUMO construct (fig. S1) by the hanging-drop vapor diffusion method at 20°C. Experiments were set up by mixing 1 μl of the protein sample [3 mg/ml in 25 mM potassium phosphate buffer (pH 7.2), 5% glycerol, and 100 μM FAD] with an equal volume of the reservoir solution containing 30% PEG4000, 100 mM sodium citrate (pH 5.6), and 200 mM ammonium acetate. Yellow crystals grew in about 4 to 5 weeks and were transferred to a cryoprotectant solution (reservoir solution with 15% glycerol) before flash-cooling in liquid nitrogen. X-ray data were collected at the X06SA beamline of the Swiss Light Synchrotron (Villigen) and at the ID14-EH1 beamline of the European Synchrotron Radiation Facility (Grenoble). Data processing and scaling were carried out with MOSFLM (30) and the CCP4 package (31) (Table 1).

The DprE1 crystal structure was solved by molecular replacement with Phaser (32) using, as a model, an ensemble constructed from six distant homologs: PDB entries 2vfr (20% sequence identity), 2i0k (15%), 3js8 (15%), 2exr (17%), 2bvf (17%), and 1w1o (17%). Selection and sequence alignment of the homologs were performed with HHPRED (33). The sequence alignment was used to trim off nonidentical side chains in Sculptor (34), and the program Ensembler ( was used to superimpose the six structures and trim nonconserved surface loops. All six individual models from the ensemble were used for automated rebuilding with phenix.mr_rosetta (35), and the best results were obtained starting from cytokinin dehydrogenase (1w1o), which gave a model with a free R factor of 33%. The model was then partly manually rebuilt in Coot (36) and refined by REFMAC5 (37). Cavity calculations were carried out with VOIDOO (38). Structural illustrations were produced with CCP4mg (39).

Crystals of full-length DprE1 (fl-DprE1, fig. S1) with BTZ043 were obtained with protein produced from the pET32b construct. The protein (about 20 μM) was incubated for 2 hours at 37°C in the presence of 25 μM FAD, 50 μM BTZ043, and 100 μM FPR in 20 mM tris (pH 7.5), 100 mM NaCl, and 1 mM MgCl2. The protein was concentrated to about 10 mg/ml on an Amicon centrifugal device (10,000 molecular weight cutoff; Millipore). Crystals were obtained by the hanging-drop vapor diffusion method at 18°C. Experiments were set up by mixing 0.5 μl of the protein sample with 1 μl of the reservoir solution containing 11% PEG300, 90 mM MES (pH 5.9), and 5% tetrabutylphosphonium bromide (Hampton Research). Yellow crystals grew in about 2 to 3 days and were transferred to a cryoprotectant (reservoir solution with 25% glycerol) before flash-cooling in liquid nitrogen. X-ray data were collected at the X06DA beamline of the Swiss Light Synchrotron (Villigen). Data processing and scaling were carried out in XDS (40) (Table 1). The DprE1-BTZ043 complex crystal structure was solved by molecular replacement with the native DprE1 structure as a model. Manual adjustments of the model were made in COOT (36), followed by refinement with REFMAC5 (37), part of the CCP4i program suite (31). In the refined structures, all residues fall in the allowed regions of the Ramachandran plot.

Subcellular localization studies with BTZ-TAMRA

BTZ-TAMRA was synthesized as described in the Supplementary Materials. Exponential cultures of M. tuberculosis grown in 7H9 medium were exposed to different concentrations of BTZ-TAMRA (1 to 12 μg/ml) for periods of 24 to 48 hours. A few microliters of culture were placed between two coverslips, and cells were observed by fluorescence microscopy with a personal Delta Vision inverted microscope (Applied Precision) equipped with a 100× objective (Olympus Plan Semi Apochromat, 1.3 numerical aperture). Cells were illuminated with Insight SSI solid-state illumination with excitation wavelengths of 475/28 [fluorescein isothiocyanate (FITC)] and 542/27 [rhodamine–Texas Red–phycoerythrin (RD-TR-PE)] and emission filters of 512/18 for FITC and 585/29 for RD-TR-PE. Exposure times varied from 0.05 to 0.2 s. Images were captured with a CoolSnap HQ2 camera and SoftWorx software (Applied Precision) and processed for publishing with Adobe Illustrator.

Supplementary Materials


Fig. S1. Sequence alignment of the DprE1 protein sequences from M. smegmatis (MSMEG_6382) and M. tuberculosis (Rv3790).

Fig. S2. SDS-PAGE showing E. coli expression of M. tuberculosis DprE1 cloned in a modified version of pET32b vector.

Fig. S3. SDS-PAGE analysis of purified M. smegmatis DprE1.

Fig. S4. Mass spectrometry analysis of the covalent DprE1-inhibitor adducts, namely, BTZ043, DNB1, and VI-9376, and a DprE1 control.

Fig. S5. Highest-scoring docked poses of DNB1 and VI-9376, following covalent docking using GOLD, in the active site of DprE1.

Fig. S6. Synthesis of BTZ-TAMRA.

References and Notes

  1. Acknowledgments: We thank V. Makarov, P. Brodin, and G. Keri for providing inhibitors; L. Menin for technical assistance with mass spectrometry; A. Liav for advice on the synthesis of FPR; K. Johnsson for comments on the manuscript; R. Orrù for technical assistance and helpful discussions; and the Protein Crystallography Core Facility of École Polytechnique Fédérale de Lausanne, the European Synchrotron Radiation Facility, and the Swiss Light Source for beam time and excellent support during x-ray data collection. Funding: Supported by the European Community’s Seventh Framework Programme (FP7/2007-2013) under grant agreement no. 260872 and by Fondazione Cariplo (no. 2008.3148). J.N. is the recipient of an International Incoming Marie Curie fellowship (252802-DPRETB) from the European Commission. F.P. is a Swiss National Science Foundation MHV Post-Doctoral Fellow. R.J.R. is funded by a Principal Research Fellowship from the Wellcome Trust (082961/Z/07/Z). Author contributions: J.N., F.P., E.M., and C.B. crystallized DprE1 and solved the structure; J.N., S.B.-R., S.B., E.F., G.D., A.P.L., and C.B. performed cloning, mutant generation, and protein expression and purification; L.R.C. performed enzymatic assays; J.N. synthesized BTZ-TAMRA; J.N. and G.Z. synthesized the FPR substrate; N.D. performed in vitro fluorescence imaging studies with M. tuberculosis; J.N., F.P., C.B., R.J.R., A.M., and D.E.E. analyzed the data; E.D.R., M.R.P., F.P., J.D.M., P.J.D., G.R., and S.T.C. supervised and directed the work; J.N., S.T.C., and C.B. wrote the paper. All authors discussed the results and commented on the manuscript. Competing interests: S.T.C. is named as inventor on the following patents or patent applications related to this work: WO2009010163; PCT-SAFE MT/FOP 20090701/; 6.1002-HU Quinoxaline derivatives and their use for the treatment of mycobacterial infections. S.T.C. is also a consultant for Alere, the company licensing this patent. The other authors declare that they have no competing interests. Data and materials availability: Coordinates and structure factors have been deposited in the PDB with access codes 4AUT for native DprE1 and 4F4Q for the DprE1-BTZ043 complex.
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