Fluorescent Benzothiazinone Analogues Efficiently and Selectively Label Dpre1 in Mycobacteria and Actinobacteria

Article abstract of DOI:10.1021/acschembio.8b00790

Benzothiazinones (BTZ) are highly potent bactericidal inhibitors of mycobacteria and the lead compound, BTZ043, and the optimized drug candidate, PBTZ169, have potential for the treatment of tuberculosis. Here, we exploited the tractability of the BTZ scaffold by attaching a range of fluorophores to the 2-substituent of the BTZ ring via short linkers. We show by means of fluorescence imaging that the most advanced derivative, JN108, is capable of efficiently labeling its target, the essential flavoenzyme DprE1, both in cell-free extracts and after purification as well as in growing cells of different actinobacterial species. DprE1 displays a polar localization in Mycobacterium tuberculosis, M. marinum, M. smegmatis, and Nocardia farcinica but not in Corynebacterium glutamicum. Finally, mutation of the cysteine residue in DprE1 in these species, to which BTZ covalently binds, abolishes completely the interaction with JN108, thereby highlighting the specificity of this fluorescent probe.

Full text of DOI:10.1021/acschembio.8b00790

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Article
Fluorescent benzothiazinone analogs efficiently and
selectively label DprE1 in mycobacteria and actinobacteria
Raphael Sommer, João Neres, Jérémie Piton, Neeraj Dhar, Astrid van der Sar, Raju Mukherjee,
Thierry Laroche, Paul J. Dyson, John D. McKinney, Wilbert Bitter, Vadim Makarov, and Stewart T. Cole
ACS Chem. Biol., Just Accepted Manuscript • DOI: 10.1021/acschembio.8b00790 • Publication Date (Web): 05 Oct 2018
Downloaded from http://pubs.acs.org on October 6, 2018
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Fluorescent benzothiazinone analogs efficiently and selectively label
DprE1 in mycobacteria and actinobacteria
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Raphael Sommer,1,§ João Neres,1,2,§,† Jérémie Piton,1,2 Neeraj Dhar,1,2 Astrid van der Sar, Raju
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Mukherjee,1,2 Thierry Laroche, Paul J. Dyson, John D. McKinney, Wilbert Bitter, Vadim
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1,2,#,*
Makarov, Stewart T. Cole
1
Global Health Institute, Ecole Polytechnique Fédérale de Lausanne, Lausanne, Switzerland;
More Medicines for Tuberculosis (MM4TB) Consortium (www.mm4tb.org);
2
3
Amsterdam UMC, Vrije Universiteit Amsterdam, De Boelelaan 1117, 1081 HV Amsterdam, The
Netherlands;
4
BioImaging and Optics Platform (BIOP), Ecole Polytechnique Fédérale de Lausanne (EPFL),
Faculty of Life Sciences (SV), Station 15, 1015, Lausanne, Switzerland;
5
Institute of Chemical Sciences and Engineering, École Polytechnique Fédérale de Lausanne,
Lausanne, Switzerland;
6
Research Center of Biotechnology of the Russian Academy of Sciences, Moscow, Russia.
§
These authors contributed equally to this work
†
Current address: UCB Biopharma, Braine l'Alleud, Belgium
# Address correspondence to Stewart T. Cole, stewart.cole@epfl.ch.
*
Current address: Institut Pasteur, 25 – 28 rue du Docteur Roux, Paris, France
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ABSTRACT
Benzothiazinones (BTZ) are highly potent, bactericidal inhibitors of mycobacteria and the lead
compound, BTZ043, and the optimized drug candidate, PBTZ169, have potential for the treatment of
tuberculosis. Here, we have exploited the tractability of the BTZ scaffold by attaching a range of
fluorophores to the 2-substituent of the BTZ ring via short linkers. We show by means of
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fluorescence imaging that the most advanced derivative, JN108, is capable of efficiently labeling its
target, the essential flavoenzyme DprE1, both in cell-free extracts and after purification, as well as in
growing cells of different actinobacterial species. DprE1 displays a polar localization in
Mycobacterium tuberculosis, Mycobacterium marinum, Mycobacterium smegmatis and Nocardia
farcinica but not in Corynebacterium glutamicum. Finally, mutation of the cysteine residue in DprE1
in these species, to which BTZ covalently binds, abolishes completely the interaction with JN108,
thereby highlighting the specificity of this fluorescent probe.
INTRODUCTION
Tuberculosis (TB), an infectious disease caused by Mycobacterium tuberculosis (Mtb), has reemerged
in recent decades as an important global health problem and was responsible for 1.8 million deaths in
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015. The main reasons for this alarming situation are the appearance of multidrug-resistant (MDR-
TB) and extensively drug-resistant (XDR-TB) strains with high mortality rates, the synergy with the
HIV/AIDS pandemic and increased poverty.2–4 The fruits of substantially increased investment in the
research on novel anti-TB drugs in the last decade led to replenishment of the drug discovery pipeline
with drug candidates in different stages of development.5,6 Among these novel, promising anti-TB
drugs are the 8-nitro-benzothiazinones (BTZs) BTZ043 with an MIC of 1 ng/mL (Figure 1) and
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PBTZ169, an improved BTZ analogue with an even lower MIC against Mtb (0.3 ng/mL) and
improved solubility, which is currently in clinical trials.7,8 The BTZs are among the most potent
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antitubercular agents known to date and act by inhibiting the enzyme decaprenyl-phospho-β-D-ribose
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oxidase (DprE1), which is one of the most attractive targets for anti-TB drug discovery. DprE1 and
DprE2 (decaprenylphosphoryl-D-2-keto erythro pentose reductase) catalyze the epimerization of
decaprenylphosphoryl-β-D-ribose (DPR) to its arabinose isomer DPA. DPA is the sole donor of
arabinose sugars that are essential for cell wall biosynthesis in Mtb.10
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BTZs are suicide inhibitors of DprE1, selectively forming irreversible covalent adducts with
Cys387 in the active site of this enzyme, as previously demonstrated by biochemical and structural
methods.11,12 A nitroso species is produced by nitroreduction and this reacts with the thiol side chain
of Cys387 to form a semi-mercaptal covalent adduct with DprE1. The nitro group of the BTZs is
crucial for their mechanism of inhibition, as its replacement by amine, hydroxylamine, pyrrole or
many other functional groups invariably led to near complete loss of antimycobacterial activity.8,12,13
We have recently shown that a fluorescently-labeled BTZ analogue (BTZ-TAMRA, Figure
1) retained modest antimycobacterial activity (MIC = 6 g/mL) and selectively labeled the poles of
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Mtb in vitro as observed by fluorescence microscopy, pointing to a polar localization of DprE1 in
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mycobacteria. Given these results we decided to investigate the SAR of the fluorescently labeled
BTZs further, in order to improve their anti-mycobacterial activity and explore their potential as
imaging agents for in vitro and in vivo studies with actinobacteria. Here, we report the development of
a highly active and selective fluorescent BTZ analogue that covalently labels DprE1, and its
application for imaging purposes. We exploit its properties to visualize the subcellular localization of
DprE1 in several BTZ-sensitive actinobacteria and track the behavior of mycobacteria in time-lapse
fluorescence microscopy.
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RESULTS AND DISCUSSION
Design and synthesis of fluorescent BTZ analogs. The first fluorescent BTZ synthesized, BTZ-
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TAMRA, showed a promising MIC of 6 g/mL against Mtb and was used as a starting point for
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further optimization. Previously reported SAR studies on the BTZ and PBTZ series of DprE1
inhibitors showed that when the 8-nitro-6-trifluoromethylbenzothiazinone core was conserved, the
moiety on position 2 could accommodate a range of substituents retaining or even improving the
7,8,14,15
MIC against Mtb.
The crystal structures of DprE1 in complex with BTZ043 and PBTZ169
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provided further insight into the SAR results, as the BTZ core is located inside the active site forming
a covalent bond with Cys387, but the 2-substituent of the BTZ ring is on the protein surface and
makes few interactions with DprE1 residues.7,11 This explains how the presence of a tri-PEG linker
between the BTZ core and the bulky fluorophore in BTZ-TAMRA provided enough space to allow
correct binding.
We designed six new fluorescent BTZ and PBTZ analogues testing different fluorophores and
spacers. The synthesis of the required tri-PEG and octyl linkers was performed through the synthetic
route shown in Scheme 1, following published procedures. 2-(2-(2-chloroethoxy)ethoxy)ethanol or 8-
chlorooctan-1-ol (3) were converted to the respective azides, which were reduced with
triphenylphosphine to the respective amines. The amines were Boc-protected and the respective
alcohol groups converted to the bromides 1 and 6. The Boc-protected linkers were then reacted with
piperazine forming the required Boc-protected piperazine-linkers 2 and 8 for subsequent coupling to
form the PBTZ-linker molecules. In addition, the Boc-protected 8-bromo-octylamine 6 was converted
to Boc-protected 8-amino-octylamine 6, in order to synthesize the alkyl linker version of BTZ-
TAMRA.
The fluorescein analog of BTZ-TAMRA was synthesized from intermediate 9 (previously
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described ) by coupling with the NHS ester of 5-carboxyfluorescein, affording JN23 (Scheme 2).
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The synthesis of the remaining analogues was started by coupling the key 3-methylthio-
benzothiazinone intermediate 1016 with each of the linkers described above, providing the Boc-
protected intermediates 11, 12 and 13. Boc hydrolysis, followed by coupling with the desired
fluorophore NHS esters in the presence of diisopropylethylamine, afforded the final compounds
JN96, JN62, JN94 and JN108. To obtain the 8-amine version of JN108, the 8-nitro group of
intermediate 13 was reduced using sodium hydrosulfite to give compound 14, which was then
subjected to Boc hydrolysis and coupling with the NHS ester of 5-carboxy-TAMRA, to afford JN105.
The propyl-TAMRA control compound JN111 was prepared by coupling n-propylamine with the
NHS ester of 5-carboxy-TAMRA.
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Anti-mycobacterial activity of novel fluorescent BTZs. The antimycobacterial potency of the
various fluorescent BTZ analogues synthesized was evaluated using the standard REMA assay (Table
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). The data shows that fluorescein-labeled JN23 was inactive against both Mtb and M. smegmatis, in
contrast to its TAMRA-labeled BTZ-TAMRA analogue. Therefore, further SAR was mostly focused
on TAMRA-labeled analogues. A BTZ-TAMRA analogue with a hydrophobic octyl linker (JN96),
prepared to assess the influence of the spacer hydrophobicity on the anti-mycobacterial activity,
presented a Mtb MIC 25-fold lower than that of BTZ-TAMRA. Similarly, the compound JN108 with
an octyl linker between a PBTZ core and TAMRA had an MIC against Mtb 400-fold lower than
JN62, the PBTZ analogue with a tri-PEG linker. JN108 also presented an MIC 4-fold lower than its
simpler BTZ analogue JN96. The smaller BODIPY-FL fluorophore was also evaluated in compound
JN94 and led to a 4-fold improvement in potency when compared to its TAMRA analogue JN62.
Compound JN105, an 8-amino version of the PBTZ-TAMRA compound JN108 was 100-fold less
potent (MIC 6.2 g/mL) than the latter, as expected (Table 1). Compounds JN105, the inactive NH2-
derivative of JN108, and JN111, a propyl-TAMRA analogue inactive against mycobacteria, were
synthesized for use as negative controls to assess the possible effect of non-specific binding. From the
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SAR studies, JN108 was the best fluorescently-labeled BTZ analogue and this was confirmed in
further work, described below.
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In vitro labeling of recombinant DprE1. Our previous study reported successful covalent labeling
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of recombinant M. smegmatis DprE1 by a BODIPY-X labeled BTZ. Using a similar strategy, we
tested the new fluorescent BTZs synthesized here for their ability to label recombinant M.
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tuberculosis DprE1, produced as described elsewhere, and analyzed the reaction products by SDS-
PAGE and fluorescence scanning. Both the fluorescein- and TAMRA-labeled BTZ analogs bound to
DprE1 irrespective of the linker used in their construction. All the nitro-BTZ fluorescent derivatives
(JN23, JN94, JN29, JN62, JN96 and JN108) bound strongly to purified DprE1 (Figure 2). JN29,
JN62, JN96, JN108 were detected in the TAMRA channel whereas JN23 and JN94 were detected in
the fluorescein channel. The non-covalent fluorescent BTZ derivative, JN105, with an amino instead
of the nitro group, bound much less strongly, as did JN111, a propyl-TAMRA derivative lacking the
BTZ pharmacophore. Similarly, very weak labeling was seen when DprE1 was inactivated by heating
for 10 min at 95°C before adding JN108 to the reaction, most likely due to non-specific binding.
Likewise, labeling was abolished when the reducing agent DTT was included. These findings are
consistent with the known mechanism of action of the BTZ12,18 and suggest that the lack of
antimycobacterial activity of several derivatives, such as JN62, may be due to its inability to reach
DprE1 in the periplasm.19
Labeling of DprE1 in M. tuberculosis cell-free extracts. To further investigate the interaction of
JN108 with its target, DprE1, and to assess the extent of binding to other proteins in M. tuberculosis,
JN108 was incubated with cell-free extracts and then analyzed by SDS-PAGE and fluorescence
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scanning. Since DprE1 is not an abundant protein, in most, but not all cases the extracts were
supplemented with purified DprE1, or with its variants, in which Cys387 had been replaced by Ser or
Gly.20 As can be seen from Figure 3, in addition to DprE1, JN108 only labeled two highly Mtb
abundant proteins with apparent molecular weights of 60 and 15 kDa. The 15 kDa band was identified
by mass spectrometry as GroES. The signal strength of the target-specific labeling was proportional to
the amount of DprE1 present and the enzyme could be detected at 0.5 M but not at 0.1 M. No
labeling occurred when BTZ-resistant DprE1 variants were used bearing C387S or C387G
substitutions and, again, this is consistent with the mechanism of action of BTZ and with prior
findings.7,8,11,20
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Exploring human hepatocyte extracts. To establish whether JN108 could label any human proteins
the compound was incubated with an extract of the human hepatocyte cell line HepG2 supplemented
with or without DprE1. From Figure 3 it is apparent that no labeling of human proteins was observed
whereas DprE1 was once again detectable at the 0.5 M level. The combined results indicate that if
off-target covalent labeling of human proteins occurs it must be below the limit of detection of this
assay.
Single cell analysis and labeling of living mycobacteria. To visualize the effect and specificity of
fluorescently labeled BTZ-derivatives on mycobacteria in real time, M. smegmatis cells constitutively
expressing GFP were grown in a microfluidic device and monitored using time-lapse microscopy.21
The culture medium was supplemented with either 700 ng/mL JN108 or its inactive NH -derivative
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(JN105, 700 ng/mL) and the results compared. When JN105 was used no growth inhibition was
observed and all bacterial cells were weakly stained on the surface although the stain could be
removed by flowing JN105-free 7H9 medium through the device (Supplementary Figure 1a,
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Supplementary Movie 2). By contrast, when JN108 was employed growth ceased within a few
hours of exposure and polar localization of the fluorophore was clearly visible in most of the cells
(Supplementary Figure 1a, Supplementary Movie 1). On continuing exposure to JN108, some
cells started to lyse (indicated by loss of GFP signal) and some cells started leaking cellular material,
which was also stained with the fluorophore.
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Similar staining was also performed using M. tuberculosis cells constitutively expressing GFP
protein. Bacteria that were grown in a microfluidic device were exposed to either JN105 or JN108
(400 ng/mL) and staining monitored by time-lapse microscopy. As was seen in the case of M.
smegmatis, the active derivative, JN108, exhibited clear polar localization with a large fraction of the
cells subsequently lysing (Supplementary Figure 1b, Supplementary Movie 3). In contrast, when
exposed to JN105, most of the cells exhibited faint staining of the whole cell surface, which was
probably indicative of non-specific binding (Supplementary Figure 1b).
Subcellular localization of DprE1. The Wag31-GFP fusion protein is an additional independent
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marker for the poles of M. smegmatis. When JN108 was used to stain DprE1 in this bacterium,
strong co-localization of the GFP and TAMRA signals could be observed at the poles using high-
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resolution microscopy (Supplementary Figure 2). When the Rv3789-GFP fusion protein was used
to label the plasma membrane of M. smegmatis or M. marinum, JN108 also displayed bright foci at
the poles of the bacilli and beyond the GFP-labeled membrane, indicating that the localization of
DprE1 is similar in M. smegmatis and M. marinum (Figure 4a and b, Supplementary Figure 3a).
Structured illumination microscopy (SIM) with JN108-stained M. smegmatis confirmed these
previous observations at the super-resolution level (Figure 4a and b).
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Labeling other actinobacteria. Many actinobacteria are susceptible to BTZ including certain
species of the genera Corynebacterium and Nocardia.8,24 This prompted us to investigate whether
DprE1 could also be localized with JN108 at the poles of species from these genera. The MIC of N.
farcinica for JN108 was evaluated at 0.01 g/mL by standard REMA techniques, confirming its
general susceptibility to benzothiazinones. On staining N. farcinica with JN108 bright foci were
visible at the poles of the cells and at the branching points, which are characteristic of Nocardia spp.,
thereby suggesting that DprE1 plays a role during tip extension not only at the poles of bacilli but also
at budding locations in branching bacteria (Figure 5a).
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The MIC of C. glutamicum for JN108 was found to be 1.1 g/mL. A mixture of M. smegmatis
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expressing a GFP-labeled membrane protein and C. glutamicum was incubated with JN108 and
examined by wide field fluorescence microscopy. In this experiment, polar labeling of M. smegmatis
was seen, as expected, but C. glutamicum cells appeared to be uniformly labeled suggesting that,
unlike mycobacteria, DprE1 is not located at the poles (Figure 5b). To determine whether this result
was a general property of Corynebacteria spp., we repeated the experiment using Corynebacterium
diphtheriae and again observed that JN108 stained cells uniformly (data not shown). Thus, for
Mycobacterium and Nocardia spp. but not for Corynebacterium spp., the localization of DprE1
indicated by JN108 staining coincides with the sites of cell wall synthesis, as can be observed by
nascent peptidoglycan staining (Supplementary Figure 4). Interestingly, when JN108 staining was
performed on non-replicating, streptomycin-starved M. tuberculosis 18b (SS18b), a model system for
latent TB,25,26 polar labeling could still be observed (Supplementary Figure 5). Since SS18b is not
susceptible to BTZ, nor to other mycobacterial cell wall inhibitors, this suggests that JN108 can still
interact with its target, DprE1, in non-replicating cells and that in growing mycobacteria the
bactericidal effect follows this interaction.
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BTZ-resistant bacteria do not display polar labeling. The mechanism of action of BTZ and PBTZ
involves the covalent binding of the drug to a conserved cysteine in the substrate-binding site of
DprE1.11,12 In vitro, mutated versions of the enzyme were not labelled by JN108 (Figure 3) and we
sought to validate that the same happens when staining living bacteria. BTZ-resistant mutants of M.
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tuberculosis and M. smegmatis were previously described. Spontaneous BTZ-resistant mutants of M.
marinum and N. farcinica were selected on 7H10 medium containing 300 ng/mL or 1 g/mL
PBTZ169, respectively. The presence of the mutations C389S and C419S in DprE1 were confirmed
by PCR sequencing of mmar5352 and nfa1970, the dprE1 orthologs in M. marinum and N. farcinica,
respectively. MICs for PBTZ169 were >1000 ng/mL and >10,000 ng/mL for resistant M. marinum
and N. farcinica, respectively, while the BTZ-sensitive parent strains had MICs of 0.3 ng/mL and
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.6 ng/mL. As expected, JN108 staining of the BTZ-resistant N. farcinica did not display polar
localization but only a faint background (Figure 6). Similarly, BTZ-resistant M. smegmatis (Figure
4c), M. marinum (Supplementary Figure 3b) and M. tuberculosis (Supplementary Figure 6) do not
display polar foci, unlike their wild-type counterparts. This confirms that labelling by JN108 in all the
tested actinobacterial species is highly specific for DprE1 and follows the same mode of action as
benzothiazinone-based antimycobacterials.
CONCLUDING REMARKS
Here, we have exploited the tractability of the 2-substituent of the BTZ and PBTZ scaffolds to attach
a range of fluorophores via an alkyl linker thereby creating tool compounds for detecting
mycobacteria in different settings. We demonstrate that the most advanced derivative, JN108, which
is TAMRA-labeled, retains significant anti-mycobacterial activity and can efficiently label its target
DprE1 in vitro, and at the poles of live mycobacteria in culture. The high specificity of JN108 is
confirmed by the absence of labeling on BTZ-resistant mutant bacteria carrying Cys→Ser mutation in
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DprE1. This exciting result shows the potential of fluorescent PBTZ analogues to detect mycobacteria
in more complex settings, such as for in vivo imaging. We are currently exploring these avenues with
JN108 in the zebrafish model of mycobacterial infection and plan to evaluate additional JN108
analogues with near-infrared range (NIR) fluorophores for their ability to detect Mtb in infected mice
by in vivo imaging techniques. Although BTZ-derived probes display weak bactericidal activity on
BTZ-sensitive bacteria, that might limit precise quantitative applications, they nonetheless have the
potential to detect M. tuberculosis in infected tissues or in sputum samples.
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METHODS
Chemistry: General Methods and Materials. All commercial reagents (Sigma-Aldrich, Acros,
Alfa-Aesar) were used as provided. All reactions were performed under an inert atmosphere of dry N2
in oven-dried (150 °C) glassware. Column chromatography was performed with a Varian 971-FP
Flash purification system with prepacked silica gel cartridges supplied by Luknova, with the indicated
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solvent system. H and C NMR spectra were recorded on Bruker Avance 400 MHz spectrometer. H
chemical shifts are reported in ppm from an internal standard of residual chloroform (7.26 ppm), or
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methanol (3.31 ppm), and C chemical shifts are reported using an internal standard of residual
chloroform (77.1 ppm), or methanol (49.0 ppm). Proton chemical data are reported as follows:
chemical shift, multiplicity (s = singlet, d = doublet, t = triplet, m = multiplet, br = broad), coupling
constant, and integration. High-resolution mass spectra were obtained on a Waters Q-TOF Ultima
instrument equipped with an ESI interface. Details of the synthetic routes used to produce the
compounds outlined in Schemes 1 and 2 are given in the Supporting Information.
Bacterial strains, cell lines, cell extracts and culture conditions. All mycobacterial strains, BTZ-
resistant mycobacterial mutants and M. tuberculosis strain H37Rv were grown at 37 °C with shaking
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in Middlebrook 7H9 (Difco) broth supplemented with 10% albumin-dextrose-catalase (ADC)
enrichment, 0.2% glycerol, 0.05% Tween 80. M. marinum strain E11 and M were grown in the same
medium at 30 °C. C. glutamicum (ATCC13032) and N. farcinica (ATCC3318) were grown in Brain-
Heart Infusion medium (Difco) at 30 °C. Plasmids based on mNeonGreen27,28 were constructed by
PCR amplification, subcloning in appropriate vectors and transformed by electroporation. To prepare
proteins, H37Rv was grown and processed as described previously.29 Briefly, cell lysates were
prepared by resuspending cell pellets in lysis buffer (PBS containing Roche protease inhibitor cocktail
tablets) and bead beating with 100 m glass beads, and were then clarified by centrifugation. The
human hepatocyte cell line HepG2 was grown in DMEM medium supplemented with Fetal Bovine
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Serum (heat inactivated). After Trypsin-EDTA treatment, ~10 cells were lysed by bead-beating and
the extract clarified by centrifugation.
Protein concentrations were determined using the
bicinchoninic acid (BCA) assay (Pierce BCA protein assay kit; Thermo Scientific).
Drug susceptibility testing. The in vitro activities against bacterial strains were measured by the
resazurin reduction microplate assay (REMA), by performing 2x serial dilutions of compounds in the
working bacterial culture, in 96 well plates (100 L final volume). Bacterial viability was determined
by adding sterile resazurin (10 L, 0.025% w/v), incubating and measuring resazurin turnover by
fluorescence (excitation/emission 560/590 nm), using a TECAN Infinite M200 microplate reader. For
M. marinum, the strains carried an integrative plasmid containing the Lux operon encoding
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luciferase and cell viability was evaluated by measuring luminescence after 3 days of exposure to
the drug.
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Fluorescent labeling of purified DprE1. Purified M. tuberculosis DprE1 (5 M) was incubated at
30 °C for 2 h with different fluorescent compounds (10 M), farnesyl-phosphoryl-beta-D-
ribofuranose (FPR; 200 M), FAD (5 M) in enzyme assay buffer (50 mM glycylglycine pH 8.0, 200
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mM potassium glutamate, 0.002% Brij-35) in a final volume of 20 L. Reactions were stopped by
addition of SDS-PAGE buffer without reducing agent and heating 5 min at 95 °C before loading on
an SDS-PAGE gel. After electrophoresis, gels were scanned using a GE Typhoon Trio™ scanner and
fluorescence was measured in the fluorescein channel for JN23 and JN94 (excitation/emission
488/520) or TAMRA channel for JN29, JN62, JN96, JN108, JN105, JN111 (excitation/emission
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23/580). Finally, gels were stained with Coomassie blue. To denature DprE1 the purified enzyme
was heated at 95 °C for 10 min before adding JN108 to the reaction.
Labeling of cell-free extracts of M. tuberculosis or hepatocytes. Cell-free extracts of M.
tuberculosis and HepG2 cells were prepared by bead-beating as described above. Proteins (68 g)
from cell-free extracts were added to the reaction mixture containing FPR (200 M), FAD (5 M),
glycylglycine (50 mM pH 8.0), potassium glutamate (200 mM), Brij-35 (0.002%) in a final volume of
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0 L. Reactions were stopped and proteins processed for fluorescence scanning as outlined above.
In some cases purified recombinant M. tuberculosis DprE1 was added to the protein extracts as a
positive control for labeling.
Time-lapse imaging. Time-lapse microscopy of bacteria was carried out as described before.21,31
Briefly, bacteria from exponentially growing cultures of M. smegmatis or M. tuberculosis were
implanted in a custom-fabricated microfluidic device31 and imaging carried out using a 100X
objective (Olympus UPLFL 100x PH, 1.3 NA) on a personal DV microscope system (GE Healthcare
Life Sciences). Images were acquired every 15 min (M. smegmatis) or every 60 min (M.
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tuberculosis), on the FITC (excitation 490/20, emission 528/38), RD-TR-PE (excitation 542/27,
emission 585/29) and phase channels. Complete 7H9 medium with or without the fluorophores
(JN108 or JN105, 400 ng/mL for M. tuberculosis, 700 ng/mL for M. smegmatis) were pumped
through the device at a flow rate of 15-20 L/min. The images were subsequently processed and
analyzed using Deltavision SoftWorx 4.1 (GE Healthcare Life Sciences) and ImageJ v 1.47a
(http://rsb.info.nih.gov/ij/).
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Subcellular localization of DprE1. To visualize JN108 labeling at the subcellular level, the bacteria
were incubated with 8 M JN108 in complete 7H9 medium, washed twice with PBS containing
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.05% Tween-80, resuspended, adsorbed on an agarose pad and covered by a glass coverslip. The
specimen was then observed with an Olympus IX81 inverted microscope or an LSM 700 inverted
confocal microscope (Zeiss). For super-resolution imaging (structured illumination microscopy), 1
L of bacterial suspension was spotted between two coverslips and imaged with a Nikon Eclipse Ti
inverted setup and images were reconstructed with NIS-Elements software.
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ASSOCIATED CONTENT
This paper contains enhanced objects and supporting information describing the methods related to
the synthesis of the different compounds and staining of nascent peptidoglycan. Supporting
information also contains supplementary figures and the description of the supplementary movies.
This material is available free of charge via the Internet at http://pubs.acs.org.
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AUTHOR INFORMATION
Corresponding author
E-mail: stewart.cole@epfl.ch
ORCID
Stewart T. Cole: 0000-0003-1400-5585
Author Contributions
J.N., R.S. N.D. and S.T.C. wrote the paper.
Notes
V.M and S.T.C. are named inventors on patents related to this work.
ACKNOWLEDGMENTS
We thank A. Vocat and S. Boy-Röttger for technical assistance, L. Vera-Cabrera for providing strains,
C. Trefzer and K. Johnsson for useful discussions. The research leading to these results received
funding from the European Community’s Seventh Framework Programme (MM4TB, Grant 260872)
and from an International Incoming Marie Curie fellowship from the European Commission to J.
Neres (DPRETB, Grant 252802). R. Sommer was supported by a grant from the Fondation
Jacqueline Beytout.
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6. Makarov, V. A. (2011) Process for the preparation of 2-amino-substituted 1,3-benzothiazine-4-
ones.
7. Neres, J., Hartkoorn, R. C., Chiarelli, L. R., Gadupudi, R., Pasca, M. R., Mori, G., Venturelli, A.,
Savina, S., Makarov, V., Kolly, G. S., Molteni, E., Binda, C., Dhar, N., Ferrari, S., Brodin, P.,
Delorme, V., Landry, V., de Jesus Lopes Ribeiro, A. L., Farina, D., Saxena, P., Pojer, F., Carta, A.,
Luciani, R., Porta, A., Zanoni, G., De Rossi, E., Costi, M. P., Riccardi, G., and Cole, S. T. (2015)
2-Carboxyquinoxalines kill mycobacterium tuberculosis through noncovalent inhibition of DprE1.
ACS Chem. Biol. 10, 705–714.
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8. Trefzer, C., Rengifo-Gonzalez, M., Hinner, M. J., Schneider, P., Makarov, V., Cole, S. T., and
Johnsson, K. (2010) Benzothiazinones: prodrugs that covalently modify the decaprenylphosphoryl-
β-D-ribose 2’-epimerase DprE1 of Mycobacterium tuberculosis. J. Am. Chem. Soc. 132, 13663–
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3665.
9. Brecik, M., Centárová, I., Mukherjee, R., Kolly, G. S., Huszár, S., Bobovská, A., Kilacsková, E.,
Mokošová, V., Svetlíková, Z., Šarkan, M., Neres, J., Korduláková, J., Cole, S. T., and Mikušová,
K. (2015) DprE1 Is a Vulnerable Tuberculosis Drug Target Due to Its Cell Wall Localization. ACS
Chem. Biol. 10, 1631–1636.
20. Foo, C. S.-Y., Lechartier, B., Kolly, G. S., Boy-Röttger, S., Neres, J., Rybniker, J., Lupien, A.,
Sala, C., Piton, J., and Cole, S. T. (2016) Characterization of DprE1-Mediated Benzothiazinone
Resistance in Mycobacterium tuberculosis. Antimicrob. Agents Chemother. 60, 6451–6459.
21. Wakamoto, Y., Dhar, N., Chait, R., Schneider, K., Signorino-Gelo, F., Leibler, S., and McKinney,
J. D. (2013) Dynamic persistence of antibiotic-stressed mycobacteria. Science 339, 91–95.
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2. Santi, I., Dhar, N., Bousbaine, D., Wakamoto, Y., and McKinney, J. D. (2013) Single-cell
dynamics of the chromosome replication and cell division cycles in mycobacteria. Nat. Commun.
4, 2470.
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3. Kolly, G. S., Mukherjee, R., Kilacsková, E., Abriata, L. A., Raccaud, M., Blaško, J., Sala, C., Dal
Peraro, M., Mikušová, K., and Cole, S. T. (2015) GtrA Protein Rv3789 Is Required for
Arabinosylation of Arabinogalactan in Mycobacterium tuberculosis. J. Bacteriol. 197, 3686–3697.
4. González-Martínez, N. A., Lozano-Garza, H. G., Castro-Garza, J., De Osio-Cortez, A., Vargas-
Villarreal, J., Cavazos-Rocha, N., Ocampo-Candiani, J., Makarov, V., Cole, S. T., and Vera-
Cabrera, L. (2015) In Vivo Activity of the Benzothiazinones PBTZ169 and BTZ043 against
Nocardia brasiliensis. PLoS Negl. Trop. Dis. 9, e0004022.
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5. Sala, C., Dhar, N., Hartkoorn, R. C., Zhang, M., Ha, Y. H., Schneider, P., and Cole, S. T. (2010)
Simple Model for Testing Drugs against Nonreplicating Mycobacterium tuberculosis. Antimicrob.
Agents Chemother. 54, 4150–4158.
6. Zhang, M., Sala, C., Hartkoorn, R. C., Dhar, N., Mendoza-Losana, A., and Cole, S. T. (2012)
Streptomycin-Starved Mycobacterium tuberculosis 18b, a Drug Discovery Tool for Latent
Tuberculosis. Antimicrob. Agents Chemother. 56, 5782–5789.
7. Hostettler, L., Grundy, L., Käser-Pébernard, S., Wicky, C., Schafer, W. R., and Glauser, D. A.
(2017) The Bright Fluorescent Protein mNeonGreen Facilitates Protein Expression Analysis In
Vivo. G3 Bethesda Md 7, 607–615.
28. Shaner, N. C., Lambert, G. G., Chammas, A., Ni, Y., Cranfill, P. J., Baird, M. A., Sell, B. R.,
Allen, J. R., Day, R. N., Israelsson, M., Davidson, M. W., and Wang, J. (2013) A bright
monomeric green fluorescent protein derived from Branchiostoma lanceolatum. Nat. Methods 10,
407–409.
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9. Chen, J. M., Boy-Röttger, S., Dhar, N., Sweeney, N., Buxton, R. S., Pojer, F., Rosenkrands, I.,
and Cole, S. T. (2012) EspD Is Critical for the Virulence-Mediating ESX-1 Secretion System in
Mycobacterium tuberculosis. J. Bacteriol. 194, 884–893.
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Balasubramanian, V., Ramachandran, V., Dhar, N., and Dinesh, N. (2014) Simple and Rapid
Method To Determine Antimycobacterial Potency of Compounds by Using Autoluminescent
Mycobacterium tuberculosis. Antimicrob. Agents Chemother. 58, 5801–5808.
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1. Dhar, N., and Manina, G. (2015) Single-cell analysis of mycobacteria using microfluidics and
time-lapse microscopy. Methods Mol. Biol. Clifton NJ 1285, 241–256.
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MIC (g/mL)
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Mtb
M. smegmatis M. marinum
2
H37Rv
mc 155
E11
ND
40
BTZ-TAMRA
JN23
6.2
>50
25
50
>50
>50
6.2
JN62
>50
ND
3.5
JN94
6.2
JN96
0.25
0.06
6.2
0.75
0.08
12.5
>50
JN108
JN105
JN111
0.1
>25
ND
>50
ND – not determined
Table 1. Minimum inhibitory concentrations of fluorescent BTZs against mycobacteria.
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LEGENDS OF THE FIGURES AND SCHEMES
Scheme 1. Synthesis of linkers: (a) Piperazine, EtOH, 60 °C; NaN , (b) DMF, 100 °C; (c) 1. PPh ,
3
3
H O, THF, rt, 2. Boc O, NaOH, ACN, H O; (d) CBr , K CO , PPh , DCM, rt; (e) NH , H O, DME,
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5 °C.
Scheme 2. Synthesis of fluorescent BTZ analogs: (a) 5-carboxyfluorescein NHS ester, DIPEA, DMF,
rt; (b) Compound 7, EtOH, 50 °C; (c) TFA, DCF, rt; (d) 5-carboxy-TAMRA NHS ester, DIPEA,
DMF, rt; (e) Compound 5, EtOH, 50 °C; (f) BODIPY-FL NHS ester, DIPEA, DMF, rt; (g)
Compound 8, EtOH, 50 °C; (h) NH Cl, NaHS, EtOH, H O, rt.
4
2
Figure 1. Structures of BTZ043, PBTZ169 and BTZ-TAMRA.
Figure 2. Labeling purified DprE1 with different fluorescent BTZ derivatives. The reactions were
performed as outlined in the Methods section and contained recombinant wild type (WT) DprE1,
boiled DprE1, DTT and the fluorescent BTZ derivatives indicated at top. After SDS-PAGE
electrophoresis the gels were Coomassie blue stained (top panel), scanned using the TAMRA channel
(Excitation 523 nm / Emission 580 nm, middle panel) of the Typhoon scanner or scanned in the
fluorescein channel (Excitation 488 nm / Emission 520 nm, lower panel). The size of the bands is
about 50 kDa.
Figure 3. Labeling cell-free extracts of M. tuberculosis (a, b) or HepG2 (c, d) cells with JN108. The
reactions were performed as outlined in the Methods section and contained JN108, JN111 or the
DprE1 enzymes, as indicated at top. After SDS-PAGE electrophoresis the gels were: (a, c)
Coomassie blue stained; (b, d) Scanned using the TAMRA channel.
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Figure 4. Super-resolution (SIM) imaging of M. smegmatis producing Rv3789-GFP (membrane,
green channel) stained with JN108 (red channel). (a) BTZ-sensitive bacteria stained with JN108 show
bright spots at their tips in the red channel. (b) Magnification of the tip on a single cell from (a)
showing JN108-stained DprE1 beyond the membrane. (c) BTZ-resistant bacteria bearing a C394S
mutation in DprE1 display only background staining when exposed to JN108 but no polar foci are
observed. Scale bars: 5 m in (a) and (c), 500 nm in (b).
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Figure 5. Labeling whole cells of N. farcinica, M. smegmatis and C. glutamicum with JN108. (a)
Staining of N. farcinica with JN108 localizes DprE1 at the poles of the cells and at nascent branching
points. Top panel, TAMRA channel fluorescence image of JN108-labeled N. farcinica. Middle panel,
corresponding phase contrast image. Lower panel, merged channels. Arrows indicate the polar
localization of the TAMRA signal and the arrowheads show foci at nascent branching points. (b)
Staining of a mixture of M. smegmatis and C. glutamicum with JN108. The top panel shows the
fluorescence image and the lower panel is the corresponding phase contrast micrograph. Arrows
indicate polar localization of DprE1 in M. smegmatis expressing GFP-labeled membrane protein
Rv3789 whereas no such polar labeling was seen with C. glutamicum. Scale bars 5 m.
Figure 6. Labeling N. farcinica carrying Rv3789-mNeonGreen as a membrane marker. (a) BTZ-
sensitive bacteria show polar DprE1 labeling (arrows). (b) BTZ-resistant bacteria do not display polar
labeling but only faint background staining. The left panel shows fluorescence in the green channel
(membrane), the middle panel shows red fluorescence (JN108) and the right panel is a merge of the
green and red channels. Scale bars, 5 m.
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Scheme 1. Synthesis of linkers: (a) Piperazine, EtOH, 60 °C; NaN , (b) DMF, 100 °C; (c) 1. PPh , H O, THF,
3
3
2
rt, 2. Boc O, NaOH, ACN, H O; (d) CBr , K CO , PPh , DCM, rt; (e) NH , H O, DME, 75 °C.
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Scheme 2. Synthesis of fluorescent BTZ analogs: (a) 5-carboxyfluorescein NHS ester, DIPEA, DMF, rt; (b)
Compound 7, EtOH, 50 °C; (c) TFA, DCF, rt; (d) 5-carboxy-TAMRA NHS ester, DIPEA, DMF, rt; (e)
Compound 5, EtOH, 50 °C; (f) BODIPY-FL NHS ester, DIPEA, DMF, rt; (g) Compound 8, EtOH, 50 °C; (h)
NH Cl, NaHS, EtOH, H O, rt.
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Figure 2. Labeling purified DprE1 with different fluorescent BTZ derivatives. The reactions were performed
as outlined in the Methods section and contained recombinant wild type (WT) DprE1, boiled DprE1, DTT and
the fluorescent BTZ derivatives indicated at top. After SDS-PAGE electrophoresis the gels were Coomassie
blue stained (top panel), scanned using the TAMRA channel (Excitation 523 nm / Emission 580 nm, middle
panel) of the Typhoon scanner or scanned in the fluorescein channel (Excitation 488 nm / Emission 520 nm,
lower panel). The size of the bands is about 50 kDa.
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