Pyrrolomycins are potent natural protonophores

Article abstract of DOI:10.1128/AAC.01450-19

The escalating burden of antibiotic drug resistance necessitates research into novel classes of antibiotics and their mechanism of action. Pyrrolomycins are a family of potent natural product antibiotics with nanomolar activity against Gram-positive bacteria, yet with an elusive mechanism of action. In this work, we dissect the apparent Gram-positive specific activity of pyrrolomycins and show that Gram-negative bacteria are equally sensitive to pyrrolomycins when drug efflux transporters are removed and that albumin in medium plays a large role in pyrrolomycin activity. The selection of resistant mutants allowed for the characterization and validation of a number of mechanisms of resistance to pyrrolomycins in both Staphylococcus aureus and an Escherichia coli ΔtolC mutant, all of which appear to affect compound penetration rather than being target associated. Imaging of the impact of pyrrolomycin on the E. coli ΔtolC mutant using scanning electron microscopy showed blebbing of the bacterial cell wall often at the site of bacterial division. Using potentiometric probes and an electrophysiological technique with an artificial bilayer lipid membrane, it was demonstrated that pyrrolomycins C and D are very potent membrane-depolarizing agents, an order of magnitude more active than conventional carbonyl cyanide m-chlorophenylhydrazone (CCCP), specifically disturbing the proton gradient and uncoupling oxidative phosphorylation via protonophoric action. This work clearly unveils the until-now-elusive mechanism of action of pyrrolomycins and explains their antibiotic activity as well as mechanisms of innate and acquired drug resistance in bacteria.

Full text of DOI:10.1128/AAC.01450-19

AAC Accepted Manuscript Posted Online 12 August 2019
Antimicrob. Agents Chemother. doi:10.1128/AAC.01450-19
Copyright © 2019 American Society for Microbiology. All Rights Reserved.
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Pyrrolomycins are Potent Natural Protonophores
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Katherine Valderrama 1
Elizabeth Pradel 1,5
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Alexander M. Firsov 2
Hervé Drobecq 1
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Hélène Bauderlique-le Roy 3
Baptiste Villemagne 4
Yuri N. Antonenko2
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Ruben Christiaan Hartkoorn 1+
1 Centre for Infection and Immunity of Lille, U1019-UMR8204, Univ. Lille, CNRS, INSERM, CHU Lille,
Institut Pasteur de Lille, Lille, France
2 Belozersky Institute of Physico-Chemical Biology, Lomonosov Moscow State University, 119991
Moscow, Russia
3 Flow Core Facility, BioImaging Center of Lille, Institut Pasteur de Lille, Lille, France
4 Drugs and Molecules for Living Systems, Univ. Lille, Inserm U1177, Institut Pasteur de Lille, Lille,
France.
5 Inserm U995 – LIRIC, Faculté de Pharmacie-Service de Bactériologie, Lille, France.
+ Corresponding author: ruben.hartkoorn@inserm.fr
Keywords:
pyrrolomycin, protonophore, uncoupler, resistance, mechanism of action, antibiotic
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27 Abstract
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The escalating burden of antibiotic drug resistance necessitates research into novel classes of
antibiotics and their mechanism of action. Pyrrolomycins are a family of potent natural product
antibiotics with nanomolar activity against Gram-positive bacteria, with a yet elusive mechanism of
action. In this work, we dissect the apparent Gram-positive specific activity of pyrrolomycins and
show that Gram-negative bacteria are equally sensitive to pyrrolomycins when drug efflux
transporters are removed, and that albumin in media plays a large role in pyrrolomycin activity.
Selection of resistant mutants allowed for the characterisation and validation of a number of
mechanisms of resistance to pyrrolomycins in both S. aureus and E. coli ΔtolC, all of which appear to
affect compound penetration, rather than being target associated. Imaging of the impact of
pyrrolomycin on E. coli ΔtolC using scanning electron microscopy, showed blebbing of the bacterial
cell wall often at the site of bacterial division. Using potentiometric probes and electrophysiological
technique with artificial bilayer lipid membrane it was demonstrated that pyrrolomycins C and D are
very potent membrane depolarising agents, an order of magnitude more active than conventional
CCCP, specifically disturbing the proton gradient and uncoupling oxidative phosphorylation via
protonophoric action. This work clearly unveils the till now elusive mechanism of action of
pyrrolomycins and explains their antibiotic activity as well as mechanisms of innate and acquired
drug resistance in bacteria.
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47 Introduction
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Recent reports have brought to light the increasingly devastating emergence of multi-drug resistant
pathogenic bacteria (1). To combat this global healthcare challenge it is essential that innovative
approaches are used to identify novel classes of antibiotics, that act through novel vulnerable
targets, and that are effective against drug resistant isolates. To date, natural products and
derivatives still account for 62% of all approved drugs and for about 80% of approved antibacterial
drugs in the period between 1981 and 2014 (2). Nonetheless, due to various factors, including
toxicity, poor pharmacokinetics and lack of in vivo efficacy, a multitude of undeveloped natural
products with potent in vitro antibiotic activity have not been further investigated. Understanding
the mechanism of action of such compounds could help evaluate their drug-like potential, and
uncover novel bacterial targets that are chemically vulnerable. To this extend this study sought to
discover the mechanism of anti-biotic activity of pyrrolomycins.
Pyrrolomycins are a group of heavily halogenated small natural antibiotics produced by
Streptomyces vitaminophilus (3), Streptomyces sp. strain UC 11065 (4) and Streptomyces fumanus
(5). Pyrrolomycins carrying a carbonyl linker between the benzyl and pyrrole moiety are reported as
potent anti-Gram-positive specific inhibitors (3, 6). Of the many natural pyrrolomycin analogues
reported, the penta-chlorinated pyrrolomycin D, has been shown to be the most potent, with an
anti-staphylococcal minimal inhibitory concentration (MIC) reported at around 1 ng/mL, and activity
against biofilms (6) though the minimal bactericidal concentration (MBC) was reported in the low
µg/mL range (7). Pyrrolomycin D cytotoxicity has been published to be moderate (6, 8), though its
acute toxicity in mice is reported at 20 mg/kg (following intraperitoneal administration) (3).
The potent activity of pyrrolomycin D could help uncover an innate vulnerability of Staphylococci,
and allow for the further chemical optimisation of this antibiotic scaffold. In this study, we present
an illuminating insight into the pyrrolomycin D spectrum of activity, describe diverse mechanisms of
bacterial resistance to pyrrolomycins and define the precise mechanism of action of these natural
products.
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77 Results:
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Chemical synthesis of pyrrolomycins C, D, I and J
Pyrrolomycins C, D, I and J (Fig. 1) were successfully produced by de novo chemical synthesis and
their structures verified by NMR (supplementary results) and high-resolution mass spectrometry.
Each of the compounds was more than 95% pure (by HPLC-UV).
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In vitro profiling of pyrrolomycin activity
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The preferential antibacterial activity of pyrrolomycins C and D against Gram-positive bacteria versus
Gram-negative bacteria was confirmed as previously published (3) (Table 1). Antibiotic susceptibility
testing in standard media (Cation Adjusted Mueller Hinton Broth, CAMHB) confirmed that both S.
aureus and S. pneumoniae were very sensitive to pyrrolomycin D, slightly less sensitive to
pyrrolomycin C, and more than 10-fold less susceptible to the methoxy analogues pyrrolomycins I
and J (Table 1). The Gram-negative bacteria tested (all cultured in CAMHB), were all an order of
magnitude less sensitive to pyrrolomycins compared to the Gram-positive bacteria, though
pyrrolomycins C and D remained more active than pyrrolomycins I and J (Table 1). Mycobacterium
tuberculosis (grown in Middlebrook 7H9 supplemented with 10% OADC, 0.2 % glycerol, and 0.05 %
Tween 80) was weakly susceptible to pyrrolomycins, with similar high MICs for all analogues (Table
1).
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To assess whether Gram-negative bacteria were resistant to pyrrolomycins due to their efficient
xenobiotic efflux systems, E. coli strains lacking a component of the tripartite AcrAB-TolC efflux
pump (E. coli ΔtolC, E. coli ΔacrA and E. coli ΔacrB), were tested for their susceptibility to
pyrrolomycins. Data (Table 1) clearly showed that the E. coli ΔtolC strain is highly sensitive to
pyrrolomycins C and D, while surprisingly the E. coli ΔacrA and E. coli ΔacrB strains remained
resistant. This suggests that wild-type E. coli resistance to pyrrolomycins is due to TolC mediated
efflux (AcrA/B independent), and not due to the absence of pyrrolomycins intracellular target. The
abundance of efficient xenobiotic efflux systems in the other Gram-negative bacteria may also
account for their insensitivity to pyrrolomycins.
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To investigate the observed insensitivity of M. tuberculosis to pyrrolomycins, experiments were
designed to evaluate the role of growth media components. Albumin (BSA) is traditionally present in
the culture media of M. tuberculosis as part of the OADC or ADC supplements, and functions to
prevent toxicity associated to the Tween 80 detergent (9). However, albumin is known to bind
drugs/compounds, and alter their free-fraction in media, potentially affecting activity. Indeed, by
using a medium supplemented with tyloxapol instead of Tween 80, it was clearly demonstrated that
the removal of BSA rendered M. tuberculosis 10 times more susceptible to pyrrolomycins (Table 1).
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In a reverse experiment, the addition of BSA or FCS (fetal calf serum) to CAMHB, reduced the activity
of pyrrolomycin against S. aureus and E. coli ΔtolC by at least 2 orders of magnitude (Table 1).
Overall, this data clearly suggests that pyrrolomycins are highly bound to albumin, and that
differences in culture media composition, particularly the presence of albumin, help explain the
previously described low sensitivity of M. tuberculosis to pyrrolomycins.
The impact of media supplementation with BSA or FCS (typically used in cell culture media) also calls
into question the validity of the previously reported high selectivity index (antibiotic activity vs
cytotoxic activity) of pyrrolomycins (6, 8). Interestingly, the cytotoxicity of pyrrolomycins observed
here against HepG2 and HEK293 human cells was in the same range as that observed for Gram-
positive bacteria grown with 10% FCS supplementation (Table 1). This data illustrates that when
correcting for differences in media composition, there is no selectivity of pyrrolomycins C, D, I or J
for bacteria versus eukaryotic cells. This finding may explain the reported acute toxicity of
pyrrolomycins in mice of 50 and 20 mg/kg for pyrrolomycins C and D, respectively (3)
Time-dependent killing and MBC.
Previous published data showed a large difference between the MIC and the minimal bactericidal
concentration (MBC) for pyrrolomycins (7). This result was confirmed here, and data shows that the
bactericidal activity is both concentration and time-dependent (Fig. S1). Following a 24-hr
pyrrolomycin D exposure, MBC (2-log drop in cfu) was achieved against S. aureus SH1000 and E. coli
∆tolC at 3 µg/mL and 0.75 µg/mL, respectively, compared to an MIC of 25 ng/mL for both strains.
Analysis of the time-dependent bactericidal activity at 200 ng/mL (8 times MIC), showed a slow but
gradual decline in cfus for both S. aureus SH1000 and E. coli ∆tolC (Fig. S1).
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Deciphering genetic resistance to pyrrolomycin D
Pyrrolomycin D resistance in E. coli ∆tolC
To gain an insight into the mechanism of action of pyrrolomycin D, efforts were made to select
resistant bacteria from E. coli ∆tolC and S. aureus streaked grown on solid media containing
pyrrolomycin D at concentrations above its MBC (1 µg/mL). Despite numerous attempts, and the
addition of up to 6x1010 bacteria per plate (100 µL of OD600=400), no resistant S. aureus clone could
be isolated. For E. coli ∆tolC, when 2 x1010 cfu were plated, two resistant colonies were isolated.
Following culture of the resistant clones in compound-free media, the resistance was confirmed to
pyrrolomycin D (MIC = 2 µg/mL). Whole genome sequencing and variant analysis of one of the
resistant mutants (E. coli ΔtolC pyrr RC1) revealed no single nucleotide polymorphism, nor short
insertion or deletion, but instead a large deletion between 4 293 549 and 4 302 404 on genome
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reference CP009273.1. This 8855-bp deletion, confirmed by Sanger sequencing, encompasses nine
genes transcribed in the order alsR, alsB, alsA, alsC, alsE, alsK, yjcS, ytcA, mdtN (Fig. 2). The second
pyrrolomycin resistant clone that was isolated contained the exact same deletion. The role of this
large deletion in pyrrolomycin resistance was confirmed using a genetic engineering approach on E.
coli strain EP664 (BW25113 ∆tolC where the kanamycin cassette was removed by FRT excision).
Replacement of the 8855 bp region in EP664 with a kanamycin resistance cassette (resulting E. coli
strain EP673) was found not to alter its pyrrolomycin D sensitivity (Table 2). However, subsequent
erasing of the kanamycin cassette by allelic exchange to generate the unmarked pyrr RC1 deletion in
strain EP676 conferred resistance to pyrrolomycin D (Table 2). This work confirmed the causality of
the 8855-bp deletion in pyrrolomycin resistance, and showed that resistance was likely due to the
new gene arrangement rather than to the gene loss.
The deletion in E. coli ΔtolC pyrr RC1 and EP676 places the mdtO and mdtP genes in close proximity
to the als operon promoter (Pals). mdtO and mdtP encode RND type multidrug efflux pump
components homologous to AcrB (inner membrane transporter) and TolC (outer membrane
channel), respectively. Thus we reasoned that the Pals promoter activity may be high in EP676 as the
local repressor gene, alsR, is truncated due to the deletion, and that mdtP overexpression could
complement the loss of tolC. To verify these hypotheses, several genetic constructs were generated
and evaluated. Firstly, when the mdtOP genes were deleted in EP676, the resulting strain, EP680,
turned pyrrolomycin sensitive, confirming the mdtOP role in pyrrolomycin resistance (Table 2).
Secondly, conditional expression of mdtP alone in EP664, a ΔtolC strain bearing an intact als-mdt
locus, lead to a pyrrolomycin resistance phenotype, showing that the overproduction of the MdtP
outer membrane channel was sufficient for pyrrolomycin resistance (Table 2). Finally, the
overexpression of alsR from a plasmid rendered EP676 sensitive to pyrrolomycin D (Table 2),
confirming the role of the als promoter and that of the AlsR repressor. Overall, these results
demonstrate that the pyrrolomycin resistance phenotype selected is due to a genetic re-
arrangement that results in the als promoter mediated upregulation of mdtP.
Selection of pyrrolomycin D resistant clones from E. coli EP680
To isolate alternative pyrrolomycin resistant E. coli mutants devoid of mutations associated with the
MdtP channel, 1 x109 EP680 (ΔtolC, Δals-mdtNOP) cells were deposited on plates containing low
concentrations of pyrrolomycin D (50 ng/mL, twice the solid MIC). Seven colonies appeared after 48
hrs. All colonies were confirmed to present low-level resistance to pyrrolomycin D (Table 3). Whole
genome sequencing and variant analysis on two selected resistant clones (EP680-RC50.6 and EP680-
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RC50.11) revealed that both isolates carried a point mutations in slyA (encoding a non-essential
transcriptional regulator of the MarR family) while the more resistant clone, EP680-RC50.6, carried
an additional point mutation in ompA (outer membrane protein A) (Table 3). These mutations were
confirmed by Sanger sequencing. The mutations in slyA were a premature stop codon at position 16
in one clone, and a point mutation at the beginning of the HTH domain (Q49R) in the other clone,
both likely generating a dysfunctional proteins. Pyrrolomycin susceptibility studies of E. coli ΔtolC
ΔslyA (EP706), confirmed that slyA inactivation confers low level pyrrolomycin resistance (Table 3).
The role of SlyA in E. coli is still poorly known, but it is a MarR-type transcriptional regulator that
targets numerous bacterial promoters (10). Though it is not clear how inactivation of slyA causes low
level pyrrolomycin D resistance in E. coli, in Salmonella enterica, slyA (91% identical) has been shown
to be involved in resistance to antimicrobial peptides (11). The mutation found in ompA (G63R) is
located in the second transmembrane ß-barrel of the protein. In contrast to this mutation, deletion
of ompA in the ΔtolC background rendered E. coli similarly sensitive (perhaps slightly more sensitive)
to pyrrolomycin D, comparable to susceptibility data reported for other antibiotics (12). While the
role of OmpA in antibiotic susceptibility is still debated, it is believed that its role in susceptibility to
non-beta-lactam antibiotics is by altering membrane integrity and permeability rather than antibiotic
transport (12), with the identified mutations presumably resulting in a gain of function phenotype.
Selection of pyrrolomycin D resistant clones from S. aureus SH1000
As no S. aureus resistant mutant could be isolated on high concentrations of pyrrolomycin D,
selection was repeated at near MIC concentrations. Three resistant colonies could be isolated when
1.5x1010 cfus were plated on 4 times MIC (100 ng/mL). As with E. coli EP680 resistant mutants, these
isolates appeared only after 48-hrs, and were confirmed to have low level of resistance to
pyrrolomycins D and C (Table 4). One of these resistant isolates (100-1) and the parental SH1000 S.
aureus strain were subjected to whole genome sequencing and variant analysis. Data showed a
point mutation in the non-essential mprF gene (L78Stop) and a second one in the region upstream of
the SAOHSC_01883 gene (conserved hypothetical), both confirmed by Sanger sequencing. Targeted
sequencing of these regions in the two other resistant isolates showed that clone 100-2 carried the
same mutations as clone 100-1, and that clone 100-3 carried a single base insertion in mprF resulting
in a premature stop codon. As different mutations in mprF were found in two resistant isolates,
mutations in this gene are likely associated with pyrrolomycin resistance.
mprF codes for
a
multiple peptide resistance factor (MprF) with phosphatidylglycerol
activity that modifies membrane phosphatidylglycerol (PG) to
lysyltransferase
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lysylphosphatidylglycerol (LPG). Increased mprF expression has been demonstrated to increase the
cationic charge of the bacterial membrane and repel cationic molecules such as daptomycin (13, 14),
methicillin (see (15) where mprF is named fmtC) and several other cationic antibacterial peptides
(16). Baring this in mind, the daptomycin susceptibility of the pyrrolomycin D resistant isolates was
determined, and all three were found to be 16-fold more sensitive compared to the parental S.
aureus strain (Table 4). This suggests that the absence of MrpF decreases the cationic charge of the
membrane and favours daptomycin entry. As pyrrolomycin D is predicted to be anionic at
physiological pH, decreasing the cationic charge of the membrane will likely disfavour its
penetration, and hence lead to low-level resistance.
Pyrrolomycin D is not an electrophile and its activity is not impacted by anti-oxidants.
Pyrrolomycin D did not react with glutathione, and is hence an unlikely electrophile that could target
proteins non-specifically (data not shown). Similarly, pyrrolomycin activity against S. aureus was not
impacted by antioxidants, such as α-tocopherol or ascorbic acid (data not shown), suggesting that
activity was not mediated through reactive radical species. To gain a first insight into whether
pyrrolomycins can affect membrane potential, the impact of pH on pyrrolomycin activity was
determined. Data clearly show that decreasing media pH improved the activity of pyrrolomycin D
against both S. aureus and E. coli ΔtolC, similarly to the effect observed with the proton uncoupler
CCCP (Fig. S2).
Visualisation of pyrrolomycin D activity by SEM
Scanning electron microscopy (SEM) was used to determine potential changes in E. coli and S. aureus
morphology following pyrrolomycin D exposure. Initial observations performed after 5 hr exposure
of 1 µg/mL of pyrrolomycin D showed that most bacteria were no longer intact. To get images of
cellular deformation prior to bacterial cell wall collapse, lower concentrations of pyrrolomycin D
were used (100 ng/mL). With respect to S. aureus, no obvious consistent bacterial deformation could
be observed following bacterial exposure. In contrast, blebbing or buldging of the E. coli ΔtolC cell
wall was clearly observed in the presence of pyrrolomycin D (Fig. 3). This deformation was largely
localised at the division septum, sometimes seen at the old pole, and was reminiscent of images of
E. coli following exposure to the antimicrobial peptide Human Defensin 5 (HD5) (17). Alltogether,
these data suggest that pyrrolomycins act by increasing the bacterial osmotic pressure that leads to
local bacterial cell wall rupture and blebs generation, or by weakening the cell wall so that it is no
longer able to contain the normal osmotic pressure.
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Impact of pyrrolomycin D on S. aureus membrane potential
To determine whether pyrrolomycins act as proton gradient depolarizing agents, the potentiometric
probe DiSC3(5) was initially used. DiSC3(5) accumulates readily in bacteria with polarised membrane,
and this strong accumulation causes self-quenching of its fluorescence signal. Upon depolarisation of
the membrane, DiSC3(5) is released, quenching is decreased and fluorescence is increased. As
previously reported (18), we found that in the absence of cells, the proton uncoupler CCCP, but also
pyrrolomycins C and D cause a concentration dependent quenching of DiSC3(5) (quenching observed
above 0.038 and 0.019 µg/mL respectively). We confirmed that this quenching is not mediated by
CCCP or pyrrolomycins absorbance of the DiSC3(5) excitation or emission wavelengths. To our
knowledge the mechanism of this interaction remains unknown. Due to the observed quenching,
compound-dependent depolarisation of bacterial membranes was evaluated at sub-quenching
concentrations. Data showed that pyrrolomycin C was particularly potent at depolarising the S.
aureus membrane, with the methylated pyrrolomycin I significantly less active. Pyrrolomycin D also
caused depolarisation at a very low concentration, however the magnitude of the depolarisation
(DiSC3(5) signal change) was inferior to that of pyrrolomycin C, but comparable to CCCP (Fig. 4).
Overall, these data suggest that pyrrolomycins C and D are more potent modulators of the
membrane potential of S. aureus than CCCP.
To confirm that pyrrolomycins cause membrane depolarisation in bacteria, an alternative
potentiometric probe, DiOC2(3), was used in combination with flow cytometry. This assay follows
DiOC2(3) changes in fluorescence from red (BL2) to green (BL1) following membrane depolarisation
(measured by BL2/BL1 ratio, Fig. 4). In contrast to DiSC3(5), we did not observe quenching of
DiOC2(3) upon pyrrolomycins or CCCP addition. In agreement with data obtained using DiSC3(5), we
found that pyrrolomycins C and D were nM modulators of membrane potential, with pyrrolomycin C
being 1 log more potent than pyrrolomycin D, and 2 orders of magnitude more potent than CCCP.
The methylated pyrrolomycins I and J were more than tenfold less potent than their respective
analogues pyrrolomycins C and D, but still able to depolarise S. aureus membrane potential at higher
concentrations (Fig. 4).
Impact on mitochondrial respiration
Mitochondrial respiration (oxygen consumption) is a direct result of the electron transport chain
maintaining of a resting proton motif force (electrochemical gradient) that drives ATP synthesis
through the ATP synthase. The addition of uncoupler depletes the proton motif force, and the
mitochondria (as well as bacteria) response by increasing proton efflux through an increase in the
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electron transport chain activity and hence respiration. In experiments measuring rat liver
mitochondrial respiration, both pyrrolomycins C and D were potent stimulators of respiration, a
feature commonly seen following the addition of protonophoric uncouplers (Fig. S3). This potent
nanomolar activity on isolated mitochondria, is likely associated with pyrrolomycin cytotoxicity to
eukaryotic cells.
Protonophore activity of pyrrolomycin D and C on a planer lipid bilayer.
To examine protonophoric activity of pyrrolomycin D and C in a pure lipid membrane system, we
measured electric current across a planar lipid bilayer (BLM) under voltage-clamp conditions (Fig.
5A). As seen from Fig. 5B, the addition of nanomolar concentrations of pyrrolomycin D to bathing
solutions at both sides of BLM induced a transmembrane electric current in a nanoampere range
(voltage was 25 mV) and the value of the current increased with pyrrolomycin D concentration.
To determine ion selectivity of pyrrolomycin D-mediated permeability, current (I) – voltage (V)
curves were measured under symmetrical (equal pH in both chambers) and asymmetrical (a
different pH in both chambers, pH1 = 7.0, pH2 = 8.2) conditions (Fig. 5C). The value of zero-current
potential (Vzero) under asymmetrical conditions was -57 mV corresponding to high proton selectivity
of the conductance (Vtheoretical = RT/F ·ΔpH = -71 mV at ΔpH = 1.2). As pyrrolomycin D, pyrrolomycin C
demonstrated good proton selectivity with a Vzero of -46 mV at ΔpH = 1.2.
To place the protonophoric activity of pyrrolomycins in the context of the conventional
protonophore CCCP, a comparison was performed at a 3-µM concentration of the compounds.
Remarkably, pyrrolomycin D demonstrated much larger protonophoric activity (BLM current) than
the uncoupler CCCP (Fig. 5C). The activity of pyrrolomycin C was lower than that of pyrrolomycin D
but still superior to that of CCCP (Fig. 5D). Larger protonophoric activity of pyrrolomycin D compared
to pyrrolomycin C can be tentatively ascribed to better permeability of the compound with five
chloride atoms (pyrrolomycin D) than with four chloride atoms (pyrrolomycin C). It has been shown
earlier that chlorination increased membrane permeability of hydrophobic anions (19, 20).
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Thanks to their many functions, natural products are an incredibly rich source of antibiotic or
antifungal molecules, but also of potent anti-cancer agents and immunomodulatory molecules.
Pyrrolomycins, produced by Streptomyces vitaminophilus and other bacteria, are described as some
of the most potent natural antibiotics known, with activities in the low nanomolar range against
Gram-positive bacteria (3), as well as activity against biofilms (6). Other natural products share a
similar chemical scaffold, such as pyoluleorin (21), TAN-876B (22) and marinopyrroles (23).
Interestingly, while all these molecules are described as antibiotics, their mechanisms of actions are
still elusive. By investigating the mechanism of action of pyrrolomycins this work highlights the
antibiotic drug potential of pyrrolomycins.
An important finding is that the antibiotic activity of pyrrolomycins is not specific to a select number
of Gram-positive bacteria, as previously reported, but is instead much more general. We clearly
demonstrated that E. coli, and probably other Gram-negative bacteria, are resistant to
pyrrolomycins due to efflux of the xenobiotic through a TolC-dependant AcrAB-independent efflux
system rather than to the absence of a target. In addition, our studies demonstrated that M.
tuberculosis insensitivity to pyrrolomycins is due to the presence of BSA in the growth media, that
likely binds pyrrolomycins and sequestrates it away from the bacteria. The impact of both BSA and
FCS in growth media calls into consideration the previously reported high selectivity window of
pyrrolomycins over eukaryotic cells in cytotoxicity assays, which contrasted with its reported acute
toxicity in mice (3). This data shows that one must be very careful when evaluating the spectrum of
activity of anti-bacterial compounds, taking great care to account for differences caused by
environmental factors such as growth media.
Investigating the mechanism of resistance to a compound can often help clarify its mode of action.
For most known antibiotics that target essential enzymes or the ribosome there is an attainable
frequency of resistance with mutations appearing in the target (rifamycins, beta-lactam antibiotics,
fluoroquinolones, aminoglycosides). However, when compounds act on non-genome encoded
targets such as membrane lipids, sugars, or peptidoglycans, as is the case for vancomycin,
teixobactin (24) or daptomycin (25), the frequency of resistance is often very low, with mutations
associated to proteins involved in compound penetration or in the formation of non-genome
encoded targets. The difficulty of obtaining target-associated mutations is suggestive of a non-
protein, non-ribosome target. The few pyrrolomycin resistant mutants that could be isolated in this
work showed low-level resistance, and the mutation most probably affects compound penetration
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by affecting membrane charge. The only mutation found to cause higher-level resistance was
identified in E. coli ΔtolC, and was due to a large genomic deletion that placed the mdtOP efflux
genes under the control of a now active promoter (due to the loss of the AlsR transcriptional
repressor). Genetic validation experiments clearly identified that overproduction of the MdtP outer
membrane channel is able to phenocopy pyrrolomycin resistance, through a mechanism that
remains to be defined, but likely equates to pyrrolomycin efflux from the bacteria, although it is
AcrAB independent.
A combination of techniques (scanning electron microscopy, pH mediated changes in MIC and
potentiometric dyes) were used to help narrow down the likely mode of action of pyrrolomycins to
the perturbation of the bacterial membrane potential. Nevertheless, membrane potential can be
perturbed by multiple mechanisms, direct or indirect (pore forming toxins, the perturbation of ion
channels, ion pumps, ionophores etc.). To this extent the BLM experiments were crucial to
demonstrate that pyrrolomycins can specifically move protons across a BLM (containing no protein)
in a dose dependant manner. This clearly defined the mechanism of action of pyrrolomycins to be
protonophores capable of depolarising bacterial membranes and uncouple their oxidative
phosphorylation. An efficient proton uncoupler needs to remain associated to the membrane, and
to have an ionisable group to allow for the transfer of protons from outside the membrane to the
cytoplasm. Both pyrrolomycins C and D are very lipophilic (predicted logP of 4.91 and 5.51
respectively, MarvinSketch 18.23.0, Chemaxon) and carry a pyrrole and a phenyl hydroxyl group that
are both ionisable groups. Analysis of pyrrolomycins I and J, suggests that the phenyl hydroxyl group
of pyrrolomycins C and D plays an important role in this proton gradient uncoupling. Predictions of
the acid dissociation constant (pKa of 5.1, Fig. S4) of the hydroxyl group of pyrrolomycins suggests it
would allow the shuttling of protons from outside to inside the bacteria, and thus uncouple the
proton motive force (as depicted in Fig. 6). The lipophilic nature of the pyrrolomycins likely favours
their localisation in the non-polar membrane lipids, cycling from the outer leaflet to the inner leaflet
to shuttle protons. To support this, the chlorine-rich, FabI-inhibitor triclosan, has also been shown to
act as a protonophore using its phenyl hydroxyl group to shuttle protons across the membrane (26).
In line with these findings, the cytotoxicity observed with pyrrolomycins is likely due to the observed
protonophore activity on mitochondrial membranes.
A number of natural products have been described to exhibit protonophoric activity, including usnic
acid (27), hypericin (28), clusianone (29), polyphenols (30) and others. Nonetheless, all these
molecules are less potent than the conventional proton uncoupler, CCCP. This work demonstrates
12

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that pyrrolomycin C and D are a magnitude more active than CCCP, and to our knowledge the most
potent known protonophores. It is plausible that natural products such as pyoluteorin, TAN-876B
and marinopyrroles, that have strong structural similarities to pyrrolomycins, may also act as
protonophores.
This work illustrates the importance of evaluating the mechanism of action of antibiotic molecules
before developing them as potential drugs. We show that pyrrolomycins are potent inhibitors of
bacterial membrane potential through their protonophore activity that allows for the specific
shuttling of protons across polarised lipid membrane bilayers. This potent activity, which is an order
of magnitude higher than the benchmark CCCP, explains its broad spectrum of antibiotic activity,
and also provides an explanation for their acute cytotoxic effects found in vitro and reported in vivo.
13

391 MATERIALS AND METHODS
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Strains and media.
Staphylococcus aureus SH1000 was kindly provided by Prof. S.J. Foster (University of Sheffield).
Escherichia coli BW25113 (CGSC#: 7636) and its derivatives E. coli ∆tolC (JW5503-1: ΔtolC732::kan,
CGSC#: 11430), E. coli ∆acrA (JW0452-3: ΔacrA748::kan, CGSC#: 11843) and E. coli ∆acrB (JW0451-2:
ΔacrB747::kan, CGSC#: 8609) were obtained from the Coli Genetic Stock Center (CGSC) and
originated from the Keio collection (31). Mycobacterium tuberculosis H37Rv was kindly provided by
the Institut Pasteur, Paris. Acinetobacter baumannii LMG1025, Pseudomonas aeruginosa LMG
12228, Klebsiella pneumoniae LMG 2095 and Streptococcus pneumoniae LMG 14545 were obtained
from the BCCM/LMG bacteria Collection. HepG2 (HB-8065) and HEK-293T (CRL-3216TM) cell lines
originated from the American Type Culture Collection (ATCC).
M. tuberculosis was conventionally grown in Middlebrook 7H9 supplemented with 10% OADC, 0.2%
glycerol and 0.05% Tween 80, or, to evaluate the impact of bovine serum albumin (BSA), in
Middlebrook 7H9, supplemented with 0.2 % glucose, 0.2 % glycerol, 0.01% tyloxapol, 0.095 % NaCl
and with or without 0.5% BSA. All other bacteria were grown in Cation Adjusted Mueller Hinton
Broth (CAMHB, Difco), alone or with supplementation of 10% fetal calf serum (FCS) or 0.5 % BSA.
Both HepG2 and HEK-293 cells were cultured and tested in DMEM supplemented with 10% FCS.
Chemical synthesis of Pyrrolomycins C, D, I and J
Pyrrolomycins C, D, I and J were chemically synthesized as described (32–34) and summarized in the
supporting information. The chemical identity and structures were confirmed by high-resolution
1
mass spectrometry and H-NMR in comparison to literature data.
Determination of MIC, MBC and cytotoxicity
The minimum inhibitory concentration (MIC) for E. coli, S. aureus, S. pneumoniae, M. tuberculosis, A.
baumannii and K. pneumonia was determined using the colorimetric resazurin microtitre assay
(REMA). Briefly, compounds dissolved in DMSO were distributed by Echo Liquid handling (Labcyte)
into the wells of a 384-well plate (500 nL of compound/DMSO per well). Plates were subsequently
incubated with a diluted bacterial culture (50 µL) and incubated (see Table S1 for conditions).
Bacterial viability was then determined by the resazurin reduction assay, as measured by
fluorescence (Ex: 530 nm, Em: 590 nm). For P. aeruginosa, that does not allow the entry of resazurin,
14

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bacterial viability was determined by bacterial optical density (OD600). For all experiments,
rifampicin or ciprofloxacin were run in parallel as a control.
Solid MIC was determined by mixing autoclaved CAMHB with 1.5 % agar with compound dilutions in
DMSO, in a 96-well plate. Bacteria were then added to the solidified media (5 µL OD=0.001), and
wells were scored for growth after 24 and 48 hr of incubation (37 °C).
The minimal bactericidal concentration (MBC) was determined by measuring the impact of
compounds on the colony forming units (cfus). Briefly, in a 96-well plate, 100 µL of bacterial
suspension was incubated in the presence of compound. The number of viable bacteria was then
determined at different time-points by plating bacterial dilutions on LB agar, and cfus determined
following overnight incubation at 37 °C.
Cytotoxicity of pyrrolomycins
For cytotoxicity assays, compounds were added in DMSO to a 384-well plate by Echo liquid handling.
50 µL of HepG2 or HEK-293 cells in DMEM (without phenol red) supplemented with 10% FCS were
then seeded into each well (5000 cells per well). Cells were incubated for 3 days (37°C, 5% CO2), and
cellular viability determined by the addition of resazurin (final concentration 0.025 %, 4 hrs), and
fluorescence determined (Ex: 530 nm and Em: 590 nm).
Isolation of pyrrolomycin D resistant mutants, whole genome sequencing and variant analysis.
The selection of E. coli or S. aureus pyrrolomycin D resistant clones was performed by plating 100 µL
of a concentrated log phase culture (concentrated to OD600 of 1, 10, 100 and 500) onto CAMHB
agar containing various concentrations of pyrrolomycin D. Plates were incubated for 48 hrs at 37 °C
to allow for the appearance of resistant colonies. Resistance of isolates was confirmed by growing
colonies on CAMHB agar without pyrrolomycin D, followed by MIC analysis. Whole genome
sequencing and variant analysis of the resistant isolates and the parental wild-type was performed
by Genoscreen (Lille, France). Identified genetic variations were confirmed by Sanger sequencing.
E. coli mutants construction
To validate that the identified alsR-mdtN deletion in E. coli ΔtolC is the cause of resistance to
pyrrolomycin D, this deletion was generated in the parental strain. For this a number of strains were
designed summarized in Fig. 2 and Table 2. The primers used are listed in Table S2.
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491
To generate EP664, the BW25113 ΔtolC::Kan Keio collection strain was transformed with pFLP2
encoding the FLP recombinase (35) to remove the kanamycin (Km) cassette. ApR transformants were
screened for Km sensitivity to assess excision of the FRT-Km-FRT cassette (Table S2). One such ApR
KmS mutant was further streaked on LB-sucrose 5% plate to cure pFLP2 and yield the unmarked
ΔtolC derivative, EP664.
EP673 was constructed from EP664 (pEP1436) by Red recombination at 30°C (36) using the PCR
product generated with primers RH296 and RH297 and the pEP1446 template (Table S2). Correct
integration of the Km1446 cassette into the als locus was assessed by PCR with primers RH278 and
RH279, and pEP1436 was cured by streaking on LB-sucrose 5% plate at 37°C.
To construct EP676, arabinose-induced EP673(pEP1436) was electro-transformed with the
hybridized 78-mer oligos RH298 and RH299 (Table S2). Clones were selected at 30°C on LB-Ap plates
containing doxycycline (100 µg/mL) and then screened for Km sensitivity. Correct allelic exchange
was confirmed by PCR with primers RH278 and RH279, and sequencing of the product. A KmS clone
was then cured of pEP1436 by streaking on LB-sucrose 5% plate at 37°C.
To obtain EP680, arabinose-induced EP676(pEP1436) was electro-transformed with the PCR product
generated with primers RH300 and RH301 on the pEP1446 template (Table S2). Integration of the
Km1446 cassette at the mdtOP locus was checked by PCR with primers RH279 and K2F. A KmR clone
was then cured of pEP1436 by streaking on LB-sucrose 5% plate at 37°C.
To construct EP705 (ompA) and EP706 (slyA), arabinose-induced EP664(pEP1436) was electro-
transformed with the PCR product generated with primers RH732 and RH733, or RH728 and RH729
(Table S2) on the pEP1446 template. Integration of the Km1446 cassette in the ompA or slyA gene
was checked by PCR with primers pairs RH734-RH735 or RH730-731, respectively. For each
construct, a KmR clone was then cured of pEP1436 by streaking on LB-sucrose 5% plate at 37°C.
Plasmids construction
To generate pEP714, mdtP was first PCR amplified from the BW25113 genome with the RH723-
RH724 primers pair and cloned into pCRbluntIITopo. The absence of mutation in the insert was
assessed by sequencing. Then mdtP was subcloned into pBAD30 as a SalI-HindIII DNA fragment.
To construct pEP712, alsR was first PCR amplified from the BW25113 genome with the RH726-
RH727 primers pair and cloned into pCRbluntIITopo. The absence of mutation in the insert was
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assessed by sequencing. Then alsR was subcloned as a SalI-XbaI DNA fragment into pFU95 digested
with the same enzymes to yield pEP712 in which alsR is transcribed from the strong constitutive
promoter PgapA. The control empty plasmid pEP713 corresponds to the intramolecular ligation of
digested pFU95.
Sample preparation for electron microscopy
Log phase E. coli ΔtolC and S. aureus SH1000 cultures grown in CAMHB were diluted to an OD600 of
0.4, and 400 µL of culture was placed in a well of a 24-well plate containing a sterile glass coverslip.
Cells were incubated for 5 hrs at 37 °C with or without 100 ng/mL of pyrrolomycin D. After
incubation, 400 µL of 2.5% w/v glutaraldehyde (in PBS) was added to the wells for 3 min. All the
supernatant was subsequently aspirated and the cells covered with 400 µL of 2.5% w/v
glutaraldehyde (overnight, 4°C). Bacteria were fixed by the addition of glutaraldehyde (2.5% w/v)
glutaraldehyde in PBS at 4 °C overnight.
Scanning electron microscopy.
After washing, the bacteria on the glass cover-slide were treated with 1 % osmium tetroxide in water
(30 min in the dark). Bacteria were then dehydrated with increasing ethanol concentration baths up
to 100% ethanol, and dried with a critical point drier (Quorum Technologies K850, Elexience,
France). The dry coverslips were then mounted on stubs and coated with 5 nm platinum (Quorum
Technologies Q150T, Elexience, France). Bacteria were observed by scanning electron microscopy
using a secondary electron detector in a Zeiss SEM Merlin Compact VP (Zeiss, France) operated at 2-
3 kV.
Membrane depolarization assay using DiSC3(5).
Experiments using 3,3-dipropylthiacarbocyanine iodide DiSC3(5), were conducted in conditions
similar to those described previously (18). Briefly, log-phase bacteria were pelleted (12000 x g, 5
min) and washed twice in buffer A (10 mM potassium phosphate buffer pH 7 containing 250 mM
sucrose) for S. aureus, and buffer B (5 mM potassium phosphate pH 7.3 and 20 mM glucose) for E.
coli ΔtolC. Bacteria were then resuspended (to OD600 0.085 for S. aureus and OD600 0.4 for E. coli
ΔtolC), in their respective buffers with 10 µM DiSC3(5) and 100 mM KCl. 100 µL of cells were added
to the wells of a 96-well plate, and placed into a pre-warmed (37°C) fluorescence plate reader
(EnVision, Perkin Elmer). Bacterial accumulation of DiSC3(5) was followed by measuring DiSC3(5)
fluorescence (Excitation 620 nm and Emission 685 nm) until it stabilized (fluorescence decreased due
to fluorescence quenching upon DiSC3(5) accumulation in the bacteria). Upon fluorescence
17

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stabilisation, different concentrations of pyrrolomycin or CCCP were added to the bacteria, and
depolarization was followed by kinetic analysis of DiSC3(5) fluorescence. Experiments were also
performed in the absence of bacteria to determine quenching of DiSC3(5) fluorescence by the
compounds themselves. Bacterial membrane depolarization was determined as the compound
mediated increase in DiSC3(5) fluorescence 10 min post exposure.
Membrane depolarization assay using DiOC2(3).
To follow bacterial membrane depolarization by 3,3'-Diethyloxacarbocyanine Iodide, DiOC2(3), S.
aureus was grown in CAMHB to a log-phase culture (OD600 ~ 0.4). Bacteria were diluted 1 in 1000 in
PBS and 1 ml added to microtubes containing 10 µL of test compounds in DMSO at 100 X final
concentration. Following a 15 min incubation at room temperature, 10 µL of 3 mM DiOC2(3) in
DMSO was added to the bacteria, mixed and incubated for a further 15 min. Bacteria were
subsequently analyzed by flow cytometry using a AttuneNxT cytometer (Thermofisher). For flow
cytometry parameters, initially the bacterial population was selected on FSC/SSC dot plot read in
logarithmic scale, using PBS as a control for background noise. Optimal parameters of threshold
were found to be FSC 2x1000 and SSC 0,1x1000. DiOC2(3) fluorescence of the bacterial population
was then analyzed in both the BL1 channel (excitation 488 nm, emission 530 nm) and BL2 channel
(excitation 488 nm, emission 590 nm) for green and red fluorescence respectively. Depolarisation
could be visualised as a decrease in BL1 fluorescence and an increase in BL2 fluorescence, and was
calculated using the BL2/BL1 ratio.
Liver mitochondria respiration assay.
Rat liver mitochondria were isolated by differential centrifugation (28) in a medium containing 250
mM sucrose, 5 mM MOPS, 1 mM EGTA, pH 7.4. The final wash was performed in the medium
additionally containing bovine serum albumin (0.1 mg/ml). Protein concentration was determined
using the Biuret method. Handling of animals and experimental procedures were conducted in
accordance with the international guidelines for animal care and use and were approved by the
Institutional Ethics Committee of A.N. Belozersky Institute of Physico-Chemical Biology at the
Moscow State University.
555
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559
Respiration of isolated mitochondria was measured using a standard polarographic technique with a
Clark-type oxygen electrode (Strathkelvin Instruments, UK) at 25°C using the 782 system software.
The incubation medium contained 250 mM sucrose, 5 mM MOPS, and 1 mM EGTA, pH 7.4. The
mitochondrial protein concentration was 0.8 mg/ml. Oxygen uptake is expressed as nmol/min·mg
protein.
18

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BLM experiments to define protonophore activity of pyrrolomycins
Bilayer lipid membrane (BLM) was formed by the brush (37) from a 2% decane solution of
diphytanoylphosphatidylcholine (DPhPC) on a 0.6-mm aperture in a Teflon septum separating the
experimental chamber into two compartments of equal 3-ml volume. Electrical current across BLM
was measured under voltage-clamp conditions with two AgCl electrodes placed into the solutions on
the two sides of the BLM via agar bridges, using a Keithley 428 amplifier (Cleveland, Ohio, USA). The
voltage applied to BLM was 50 mV.
Acknowledgement
This work was funded by the ATIP Avenir young investigator program (CNRS/INSERM). We are
thankful for the use of the NMR facility of the Institut Pasteur de Lille, a facility co-funded by the
European Union with the European Regional Development Fund (ERDF), by the Hauts-de-France
Regional Council (contrat n°17003781), Métropole Européenne de Lille (contract n°2016_ESR_05),
and French State (contract n°2017-R3-CTRL-Phase 1). We thank Nicolas Barois (CIIL) and the
BioImaging Center Lille for access to the SEM, equipment supported by the ANR (10-EQPX-04-01)
and the EU-FEDER (12,001,407). We thank Alexandre Vandeputte and the HCS facility « Ariadne »
that was supported by ANR and by the Feder (ANR-10-EQPX-04-01 / Equipex Imaginex BioMed). We
thank the Flow Core Facility – BioImaging Center Lille for access to AttuneNxT cytometry.
19

581 Tables
582
583
584
585
586
Table 1. Minimum inhibitory concentrations (MIC95) of pyrrolomycins C, D, I and J on a panel of bacteria in
indicated growth media. In addition the cell line cytotoxicity is indicated for the pyrrolomycins as IC95 [IC50].
Data represents the average of at least 3 replicates. Rifampicin (Rif) and ciprofloxacin (Cipr) were used as anti-
microbial controls.
MIC95 (µg/mL)
Strains
Culture media
Pyr D
Pyr C
Pyr J
Pyr I
Rif
Cipr
S. aureus SH1000
S. pneumoniae
CAMHB
CAMHB
CAMHB
CAMHB
CAMHB
CAMHB
CAMHB
CAMHB
CAMHB
0.025
0.1
0.1
6.25
27
1.5
0.4
>50
>50
>50
>50
1
3
0.05
0.05
>0.2
>0.2
>0.2
>0.2
>0.2
/
1
0.006
3
1
A. baumannii
3
>50
>50
>50
>50
1.5
/
0.2
K. pneumoniae
5
0.03
0.125
0.015
<0.015
<0.015
<0.015
P. aeroginosa POA1
E. coli BW25113
12.5
33
3
6
E. coli BW25113 ∆tolC
E. coli BW25113 ∆acrA
E. coli BW25113 ∆acrB
0.025
0.0125
/
3
3
/
/
/
/
/
M. tuberculosis H37Rv
7H9-GT-OADC
3
3
4
6
0.005
0.5
S. aureus SH1000
S. aureus SH1000
CAMHB +BSA
CAMHB +FCS
25
21
12.5
21
>50
>50
>50
>50
0.05
0.05
1
1
E. coli BW25113 ∆tolC
E. coli BW25113 ∆tolC
CAMHB +BSA
CAMHB +FCS
33
6
50
25
25
17
>0.2
>0.2
0.015
12.5
1.5
<0.015
M. tuberculosis H37Rv
M. tuberculosis H37Rv
7H9-GTy-ND
0.2
3
0.13
1.5
0.83
25
0.4
25
0.02
0.1
0.25
0.5
7H9-GTy-ND+BSA
HepG2 (IC95 [IC50])
HEK-293
DMEM+FCS
DMEM+FCS
3 [1.1]
1.5 [0.62]
3 [1.01]
25 [8.3]
29 [8.6]
3 [1.00]
25 [8.52]
25 [7.19]
587
588
589
590
591
592
593
594
595
CAMHB = cation adjusted Mueller-Hinton Broth
CAMHB+BSA = cation adjusted Mueller-Hinton Broth supplemented with 0.5% (w/v) BSA fraction V
CAMHB+FCS = cation adjusted Mueller-Hinton Broth supplemented with 10% (v/v) fetal calf serum
7H9-GT-OADC = Middlebrook 7H9 media supplemented with 0.2 % glycerol, 0.05% Tween 80 and 10% OADC (final concentrations: 0.005%
oleic acid, 0.5% albumin fraction V, 0.2% dextrose, 4 mg/L catalase and 0.95 g/L NaCl)
7H9-GTy-ND = Middlebrook 7H9 media supplemented with 0.2 % glycerol, 0.025% tyloxapol, 0.2% dextrose and 0.95 g/L NaCl
7H9-GTy-ND+BSA = Middlebrook 7H9 media supplemented with 0.2 % glycerol, 0.025% tyloxapol, 0.2% dextrose and 0.95 g/L NaCl and
0.5% albumin fraction V
DMEM+FCS = Dulbecco's Modified Eagle Medium supplemented with 10% (v/v) fetal calf serum
20

596
Table 2: Resistance profiles of E. coli BW25113 ΔtolC strains to pyrrolomycins D and C
MIC95 (µg/mL)
Strain
Characteristics
Pyr D
Pyr C
Km
ΔtolC
ΔtolC::Km
0.025
3
0.125
6
>50
>50
4
ΔtolC pyrr RC1
EP664
ΔtolC::Km, with selected pyrrolomycin resistance
ΔtolC
0.025
0.025
3
0.125
0.125
6
EP673
ΔtolC ΔalsRBACEK-yjcS-ytcA-mdtN::Km
ΔtolC ΔalsRBACEK-yjcS-ytcA-mdtN
ΔtolC ΔalsRBACEK-yjcS-ytcA-mdtNOP::Km
empty plasmid bearing PgapA (control)
PgapA-alsR plasmid (alsR overexpression)
empty plasmid w/o arabinose
>50
4
EP676
EP680
0.025
0.5
0.125
>50
EP676 (pEP713)
EP676 (pEP712)
EP664 (pBAD30)
EP664 (pBAD30)
EP664 (pEP714)
EP664 (pEP714)
0.062
0.062
0.062
0.062
1
empty plasmid with 10mM arabinose
pEP714 is pBAD30::mdtP, without arabinose
pEP714 is pBAD30::mdtP, with 10mM arabinose
597
598
599
Table 3: Pyrrolomycin D sensitivity of E. coli tolC strains on solid media
E. coli strain
Characteristics
Solid pyrrolomycin
D MIC (ng/mL)
EP680
ΔtolC ΔalsRBACEK-yjcS-ytcA- 40
mdtNOP::Km
EP680-RC:50.6
EP680-RC:50.11
EP664
slyA (Q49H), ompA (G63R)
slyA (W16stop)
ΔtolC
80
60
40
30
50
EP705
EP664 ΔompA
EP664 ΔslyA
EP706
600
601
602
603
604
Table 4: Sensitivity of S. aureus SH1000, and resistant isolates to pyrrolomycin D and daptomycin as measured
on solid media.. The genetic variations identified by whole genome sequencing (WGS) or targeted Sanger
sequencing (S) are shown.
S. aureus strain
SNP identified relative to SH1000
Solid pyrrolomycin D
Daptomycin MIC95
(µg/mL)
MIC (ng/mL)
SH1000
100-1
WT
20
35
1.25
mprF (L78Stop) (WGS) (S)
1797388 (t->g) (WGS) (S)
mprF (L78Stop) (S)
1797388 (t->g) (S)
0.078
100-2
30
0.078
21

100-3
mprF (Y419Stop) (S)
35
0.078
605
22

606 Figures
607
Pyrrolomycin R1 R2
C
D
I
H
Cl
H
H
H
CH3
J
Cl CH3
608
609
Fig. 1. Chemical structures of pyrrolomycins C, D, I and J
610
23

PalsR
4293549
|
4302404
|
ytcA
E. coli ΔtolC
fdhF
mdtP mdtO mdtN
yjcS
alsK
rpiB
alsE
alsC
rpiB
alsC
alsA
alsB
alsR
|
8855 kb
|
|
<
<
<
<
<
mdtN alsR
E. coli ΔtolC pyrr RC1
<
fdhF
ytcA
mdtP mdtO
<
<
X
PalsR
4293549
|
4302404
|
fdhF
mdtP mdtO mdtN
yjcS
alsK
alsE
alsA
alsB
alsR
rpiB
EP664
|
8855 kb
|
|
|
|
<
<
<
<
<
<
<
<
<
<
mdtN
(Km cassette 1446)
<
<
<
alsR
fdhF
mdtP mdtO
<
<
<
rpiB
EP673
|
|
<
<
<
<
<
mdtN alsR
<
rpiB
rpiB
fdhF
mdtP mdtO
EP676
<
<
<
<
<
<
<
<
<
<
<
<
<
<
<
<
<
Km cassette
1446)
<
<
<
<
P
<
N
alsR
fdhF
<
<
<
EP680
<
611
612
613
614
615
616
617
618
619
620
621
622
623
Fig. 2. Illustration of the genetic arrangement found in pyrrolomycin resistant E. coli isolates, and engineered
validation strains. From the parental E. coli ΔtolC::Km strain, a pyrrolomycin D resistant isolate E. coli ΔtolC
pyrr RC1 was sequenced and shown to bear a 8855-bp deletion between base 4293549 and 4302404, deleting
7 whole genes and 2 partial genes, and forming a hybrid in-frame gene between the truncated alsR' and 'mdtN
genes. The disruption of the alsR gene (encoding a transcriptional repressor), likely leads to de-repression of
the als promoter Pals. Deletion mutants were generated via Red-recombineering to validate that resistance was
due to mdtOP expression via Pals de-repression. To achieve this, the original E. coli ΔtolC::Km strain was deleted
of its Km cassette to obtain EP664 ΔtolC. The 8855-bp als locus was then deleted and replaced with a Km
cassette transcribed in the orientation opposite the als and mdt genes, to yield EP673. EP673 was then cured
of the Km cassette to generate EP676. Finally, to determine the involvement of mdtO and mdtP, these genes
were deleted and replaced by a Km cassette to obtain EP680.
624
24

A
B
C
D
625
626
Fig. 3: SEM images of E. coli ΔtolC following a 5-hrs exposure to DMSO (A) or 100 ng/mL of pyrrolomycin D (B-
627
628
D). Cell wall blebs can be clearly observed on pyrrolomycin exposed bacteria (indicated by white triangles).
629
25

A
B
10000
8000
6000
4000
2000
0
4
3
2
1
0
*
*
*
*
*
0.1
1
10
100
1000
10000
1
10
100
1000
[Compound] (nM)
[Compound] (nM)
630
631
Fig. 4: S. aureus membrane potential measured using potentiometric dyes DiSC3(5) (A) and DiOC2(3) (B) in the
presence of pyrrolomycin C (filled circle, solid line), D (filled square, solid line), I (filled triangle, , dotted line)
J (filled triangle, , dotted line) and CCCP (open circle, solid line). Asterisks indicate concentrations at which
the molecules quench the fluorescence signal of DiSC3(5).
632
633
634
635
26

636
637
638
Fig. 5. (A). A schematic of the BLM (made from DPhPC) electrophysiological setup used to determine
protonophoric activity of pyrrolomycin. (B) Induced electrical current through planar BLM following
the addition of indicated concentrations of pyrrolomycin D (pyrD) to both the cis and trans
chambers. The solution was 50 mM Tris, 50 mM MES, 10 mM KCl, pH 7.0. BLM voltage 25 mV. (C)
Current-voltage (I-V) curves for 1 µM pyrrolomycin D under symmetrical (Both cis and trans
639
640
641
642
27

643
644
645
646
chambers at pH 7.0, black line, or both at pH 8.2, red line) and asymmetrical conditions (cis chamber
pH 7.0, trans chamber pH 8.2, blue line). (D) Electrical current (nA) induced through DPhPC
membrane by 3 µM of pyrrolomycin D, pyrrolomycin C and CCCP (Mean±S.D., n=4).
28

647
648
649
650
Fig. 6. Model of the proton gradient uncoupling by pyrrolomycins. Lipophilic pyrrolomycins likely accumulate
in the non-polar lipid membrane because of their physical chemical properties. When oriented to the less
acidic internal side of the cytoplasmic membrane leaflet, the pyrrolomycin hydroxyl group will preferentially
ionize, releasing a proton into the cytoplasm. When flipped to the outer leaflet of the membrane, the more
acidic environment will favour the protonation of the hydroxyl group, and hence the uptake of an extra-
cytoplasmic proton, which in turn will be released in the cytosol. Overall, the cycling of this process will allow
for the net movement of protons from outside the bacterial cell to the inside, uncoupling the proton gradient,
and disturbing the membrane proton motive force.
651
652
653
654
655
656
657
658
659
29

660 References
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663
664
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669
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673
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693
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