Bactericidal urea crown ethers target phosphatidylethanolamine membrane lipids

Article abstract of DOI:10.1039/d1ob00263e

An increasing number of people are infected with antibiotic-resistant bacteria each year, sometimes with fatal consequences. In this manuscript, we report a novel urea-functionalized crown ether that can bind to the bacterial lipid phosphatidylethanolamine (PE), facilitate PE flip-flop and displays antibacterial activity against the Gram-positive bacterium Bacillus cereus with a minimum inhibitory concentration comparable to that of the known PE-targeting lantibiotic duramycin. This journal is

Full text of DOI:10.1039/d1ob00263e

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Bactericidal urea crown ethers target
phosphatidylethanolamine membrane lipids†
Cite this: Org. Biomol. Chem., 2021,
19, 3838
Sarah R. Herschede,a Hassan Gneid, a Taylor Dent,a Ellen B. Jaeger,a
Received 11th February 2021,
Accepted 8th April 2021
a
Louise B. Lawson b and Nathalie Busschaert
*
DOI: 10.1039/d1ob00263e
rsc.li/obc
An increasing number of people are infected with antibiotic-resist- bacterial cells, which display larger amounts of negatively
ant bacteria each year, sometimes with fatal consequences. In this charged phospholipids than mammalian cells.4,5 However, the
manuscript, we report a novel urea-functionalized crown ether use of AMPs for the systemic treatment of bacterial infections
that can bind to the bacterial lipid phosphatidylethanolamine (PE), is hindered by their poor pharmacokinetics, high production
facilitate PE flip-flop and displays antibacterial activity against the cost, high dosage requirements, and risk for resistance due to
Gram-positive bacterium Bacillus cereus with a minimum inhibi- proteases.6 It is therefore beneficial to develop non-peptidic
tory concentration comparable to that of the known PE-targeting molecules that can bind to bacterial lipids and exert antibac-
lantibiotic duramycin.
terial activity in a similar fashion to AMPs.
Supramolecular chemists have started to develop small mole-
cules that bind to lipid headgroups, but the focus has been on
mammalian lipids such as phosphatidylserine (PS),7–10 phos-
phoinositides (PI),11,12 and phosphatidylcholine (PC).13,14
Where bacterial membranes have been targeted, it has
been limited to polycationic species that bind to the
negatively charged lipids in bacteria (e.g., cardiolipin and
phosphatidylglycerol).15–24 However, the extensive use of non-
selective coulombic interactions creates a risk for off-target
effects and human toxicity. In contrast, here we describe the
development of neutral non-peptidic compounds that target the
bacterial lipid phosphatidylethanolamine (PE). PE is a zwitter-
ionic phospholipid found in most Gram-negative bacteria25,26
and in certain Gram-positive bacteria.5 In mammalian cells, the
most common zwitterionic lipid is phosphatidylcholine (PC).4
PE and PC have a similar headgroup but differ in their degree
of methylation of the ammonium group (Scheme 1a), allowing
selective binding of PE over PC. Two AMPs, duramycin and cin-
namycin, have been shown to selectively target PE in bacterial
membranes and apoptotic cells.27–29 Yet, the number of non-
peptidic small molecules that target PE is very limited,30–37 and
it is thus desirable to develop novel small molecules with high
PE selectivity and antibacterial potency.
Infectious diseases represent a leading cause of death world-
wide. While the advent of antibacterial agents has led to much
improvement, most antibiotics in clinical use today were devel-
oped during the 1940s to 1960s.1 This lack of novel drugs has
given bacteria time to develop numerous resistance mecha-
nisms against the most commonly used antibiotics. There is
thus an urgent need for the development of new antibiotics
with a low chance of inducing resistance. One drug target that
has become increasingly popular in this regard is the bacterial
membrane. It contains one third of the proteins in the bacter-
ium and is the site for crucial biological processes which could
be disrupted with membrane-binding antibiotics.2 Resistance
is thought to be less likely due to the rapid bactericidal effect
of membrane disruption, and the fact that lipid mutations are
less trivial than protein mutations. Unsurprisingly, there are
many natural products with antibacterial activity that function
by targeting the membrane, most notably antimicrobial pep-
tides (AMPs).3 AMPs are a large family of naturally occurring
peptides that are usually poly-cationic and amphiphilic in
nature.3 It is generally believed that the cationic charge is
responsible for membrane binding and the selectivity towards
Our approach in developing a PE targeting receptor utilizes
a urea functionality to bind to the phosphate moiety in PE,38
18-crown-6 to bind to the ammonium group of PE,39 and a tri-
fluoromethyl-substituted phenyl substituent as a lipophilic
membrane anchor40 (Scheme 1b). The urea and crown ether
functionality are linked together through either a rigid linker
(1a) or a flexible linker (1b) to determine the geometry that
aDepartment of Chemistry, Tulane University, New Orleans, Louisiana 70118, USA.
E-mail: nbusschaert@tulane.edu
bDepartment of Microbiology and Immunology, Tulane University School of
Medicine, New Orleans, Louisiana 70112, USA
†Electronic supplementary information (ESI) available: Synthesis and character-
ization of the compounds, experimental details for NMR and fluorescence titra-
tions, liposome studies and bacterial studies. See DOI: 10.1039/d1ob00263e
3838 | Org. Biomol. Chem., 2021, 19, 3838–3843
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Table 1 Overview of the PE-binding and antibacterial ability of hosts
1a–2b and 18-crown-6. All data is the average of at least 3 independent
repeats and errors represent standard deviations
Ka (M−1), NMRa
Host POPE POPC
KSV (M−1), fluorescenceb
NBD-PE NBD-PC
MICc
B. cereus
(μM)
1a
1b
2a
2b
531 56 72
6
(6.3 0.8) × 104 (1.3 0.1) × 104 25–30
Weakd
263 18 129
Weakd
22
Weakd Weake
Weake
n.d. f
>100
>100
6.25
9
3
n.d. f
n.d. f
n.d. f
18C6 180 45 n.d.g
Weake
Weake
>100
a Association constant (Ka, M−1) obtained through 1H NMR titrations
in 0.5% H2O : 24.5% DMSO-d6 : 75% CDCl3 at 298 K. b Stern–Volmer
constant (KSV, M−1) obtained through titrations of the hosts into POPC
or 1 : 1 POPE : POPC liposomes containing NBD-labelled lipids.
c Minimum inhibitory concentration (MIC) obtained using broth
microdilution methods. d No significant change in chemical shift was
observed. e No significant change in fluorescence intensity was
observed. f Not determined (n.d.) due to insolubility. g Not determined
(n.d.) due to lack of protons capable of H-bonding.
Scheme 1 (a) Structure of the head group of the targeted bacterial
lipid PE and the mammalian lipid PC. (b) Structure of the putative PE-
binding compounds 1a and 1b and the proposed complex of 1a with PE.
(c) Structure of control compounds 2a and 2b.
measurable interactions with either lipid. Computational mod-
elling using Molecular Operating Environment (MOE) suggests
that the flexible linker allows an intramolecular H-bond
between the urea NHs and crown ether oxygens, thereby block-
ing the binding site (ESI†). The control compounds 2a, 2b and
allows the best binding to the head group. Control compounds 18C6 did not bind as effectively as 1a either. The “rigid
2a, 2b, and 18-crown-6 (18C6) were also synthesized to investi- control” 2a showed non-selective binding to both lipids, while
gate the importance of the urea or crown ether moieties in the the “flexible control” 2b only showed minimal binding to
molecular recognition of PE lipids (Scheme 1c). Synthetic POPC (Table 1). The stronger interaction of POPE with 2a
details and characterization are provided in the ESI.† Crown versus 2b is probably due to hydrogen bonding between the
ether derivatives have previously been reported as anti- methoxy substituents of 2a and the ammonium group of
microbial agents due to their ability to function as ionophores POPE. Association constants with 18-crown-6 could only be
for K+ ions.41 However, the low membrane selectivity of iono- determined for POPE due to the lack of protons capable of
phores has impaired their clinical usefulness in most cases. In H-bonding in both POPC and 18C6. However, binding studies
contrast, our design takes advantage of 18C6’s known ability clearly showed that the crown ether was able to complex POPE
to selectively bind to primary ammonium cations over more (Ka = 180 M−1), but to a smaller extent than compound 1a
substituted ammonium cations.39 Combined with the phos- which can coordinate both the ammonium and phosphate
phate binding unit and membrane anchor, the crown ether groups of POPE.
derivatives become PE-selective membrane-active agents with
improved antibacterial potency.
After observing selective binding in organic solvents, we
investigated the interaction of the compounds with lipids that
To assess the selectivity of 1a and 1b for PE over PC lipids, are part of phospholipid membranes using fluorescence titra-
we initially performed a set of 1H NMR titrations in organic tions. Due to the inability of pure POPE to form stable lipo-
solvents. Under these conditions the lipids do not form mem- somes,43 the titrations were performed with either POPC lipo-
branes but are free in solution. While this is not a perfect somes containing
1 mol% 18:1-06:0 NBD-PC, or 1 : 1
mimic of biological conditions, it allows an accurate determi- POPE : POPC liposomes containing 1 mol% 18:1-06:0 NBD-PE
nation of association constants and a good indication of the (ESI†). The NBD fluorophore in the labelled lipids is attached
inherent head group selectivity of each compound. The titra- to the acyl chain of the lipids, but is known to loop up to the
tions were performed in 0.5% Milli-Q H2O, 24.5% DMSO-d6 polar membrane–water interface.44 As such, NBD-labelled
and 75% CDCl3 for solubility reasons and using either POPE lipids function as surface probes and have been used to
(1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine)
or monitor lipid phase separation,45 and lipid binding and
POPC (1-palmitoyl-2-oleoyl-glycero-3-phosphocholine) as guest partitioning.46–48 The addition of 1a to PE-containing lipo-
(ESI†). The data was fitted using Bindfit42 and the obtained somes caused complete quenching of the NBD fluorophore
association constants (Ka) are given in Table 1. The 1H NMR (Fig. 1). Fluorescence quenching was less pronounced when 1a
titrations confirmed that rigid compound 1a binds more was added to PC liposomes, suggesting selective binding of PE
strongly to POPE (Ka = 531 M−1) than POPC (Ka = 72 M−1). over PC. Quenching of the NBD-labelled lipids by compound
Surprisingly, the flexible crown ether analog 1b showed no 1a showed a Stern–Volmer relationship (Fig. 1, inset), and the
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Fig. 1 Binding of 1a to liposomes. Fluorescence spectra obtained
through the titration of 1a into a solution of 1 : 1 POPC : POPE liposomes
containing 1 mol% NBD-labelled PE. Spectra were normalized based on
the maximum intensity of the spectrum before the addition of 1a. The
inset shows the Stern–Volmer plots of the fluorescence titrations of
compound 1a into either 1 : 1 POPC : POPE liposomes containing 1 mol%
NBD-labelled PE (‘PE’ green squares) or pure POPC liposomes contain-
ing 1 mol% NBD-labelled PC (‘PC’, purple open circles). Plots are the
average of at least 4 independent Stern–Volmer plots, and error bars
represent standard deviations.
Fig. 2 Lipid flip-flop induced by 1a–2b. (Top) Experimental set-up:
100 nm DOPC liposomes (25 µM) containing fluorescent NBD-PE in the
outer leaflet of the membrane are incubated with 25 µM 1a–2b,
18-crown-6 or DMSO to induce lipid flip-flop. At certain time intervals,
the fluorescence of the NBD-PE lipids in the outer leaflet is quenched
by the addition of dithionite to calculate the % of NBD-PE that has been
flipped by the compounds. (Bottom) Percent of NBD-PE flipped by
25 µM 1a–2b, 18-crown-6 or DMSO over a time scale of 60 minutes.
Plots are the average of at least 4 independent repeats, and error bars
represent standard deviations.
obtained Stern–Volmer constants further confirmed the
selectivity of 1a for PE over PC (Table 1). None of the other
compounds tested showed any fluorescence quenching with
either PE or PC lipids (ESI†). Overall, this data supports our
previous finding that 1a can bind strongly and selectively
to PE.
stronger PE binding ability for 1a observed in the 1H NMR and
To provide further evidence of favourable binding to PE in fluorescence titrations. In fact, facilitated PE flip-flop could be
lipid bilayers, we performed other liposome-based experiments observed for 1a at concentrations as low as 3.125 µM (ESI†),
such as calcein leakage assays49 and lipid flip-flop assays.50 which is a significant improvement on a previously reported
The calcein leakage assays did not indicate membrane disrup- synthetic crown ether sulfonamide that could only mediate
tion or pore formation by any of the crown ether derivatives modest PE flip-flop at high concentrations (100 µM).52 Flip-
(ESI†). In contrast, the lipid flip-flop studies showed more flop of PC lipids was not observed for any of the compounds,
promising results. In this assay, 100 nm unilamellar DOPC further confirming the high selectivity of 1a for PE over PC
(1,2-dioleoyl-sn-glycero-3-phosphocholine) liposomes were pre- (ESI†).
pared containing 1 mol% NBD-PE or NBD-PC in the outer
Lastly, we wanted to determine if the PE-targeting com-
leaflet of the membrane. Translocation or ‘flip-flop’ of phos- pounds possess antibacterial activity. PE is found in the inner
pholipids across a lipid bilayer is normally a very slow process membrane and the inner leaflet of the outer membrane of
with a half-life of a few hours.51 In the presence of molecules Gram-negative bacteria, rendering access to PE in Gram-nega-
that can bind to the lipid head group, the polarity of the head tive bacteria challenging.25,26 On the other hand, most Gram-
group can be reduced and lipid translocation can be positive bacteria lack PE, except for species of Bacillus and
facilitated.50,52 To detect flip-flop, the NBD group of lipids in Clostridium. Any compound targeting PE is therefore expected
the outer leaflet can be selectively quenched via reduction with to function as a narrow-spectrum antibacterial agent against
membrane-impermeable dithionite.53 Residual fluorescence these bacterial species. Narrow-spectrum antibiotics are
will be the result of flip-flop of the NBD-labelled lipid from the gaining popularity because they do not select for resistance in
outer leaflet of the membrane to the inner leaflet. The results non-pathogenic bacteria and do not impact the human micro-
for the PE flip-flop assay are given in Fig. 2. Only compound 1a biome.54 With this in mind, we performed a screening assay
is able to facilitate PE translocation, in agreement with the where the compounds were incorporated into a Müller-Hinton
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agar medium, and the agar was subsequently inoculated with
the bacterial species S. simulans (0% PE),55 B. subtilis (20–30%
PE),56 and B. cereus (40–50% PE)57 (ESI†). Compounds 1b, 2a
and 18-crown-6 did not inhibit the growth of any of the bac-
teria, consistent with their lack of activity in the assays
described above. In contrast, 2b showed antibacterial activity
against all bacteria tested, regardless of their PE content. This
indicates that 2b exerts its antibacterial activity through a
mechanism that does not involve PE binding, consistent with
1
its lack of PE binding observed in the H NMR titrations and
flip-flop assays. More interestingly, compound 1a had no effect
on the growth of S. simulans, caused a significant delay in the
growth of B. subtilis and complete inhibition of bacterial
growth of B. cereus. The correlation with the PE-content of
these bacterial species suggests that the mechanism of 1a
involves binding to PE lipids.
The antibacterial activity of 1a against B. cereus was sub-
sequently investigated in more detail. B. cereus is a common
cause of foodborne illness,58 is closely related to the bioter-
rorism agent B. anthracis59 and is therefore a pathogen of
interest. The minimum inhibitory concentration (MIC) of
compounds 1a–2b against B. cereus was determined using
standard broth microdilution methods60 (Table 1 and ESI†).
Compound 1a showed an MIC value of 25–30 µM, compar-
able to the MIC value obtained for the known PE-targeting
peptide duramycin (MIC ∼32 µM, ESI†). Membrane-active
antibiotics
are
normally
bactericidal
rather
than
Fig. 3 Membrane depolarization of B. cereus by 1a measured using
Disc3(5). (a) Fraction of Disc3(5) released after 15 minutes incubation
with clindamycin (negative control, 1 µg mL−1, 1× MIC), 25 µM 1a (1×
MIC), 40 µM 1a (1.6× MIC), 250 µM 1a (10× MIC) or gramicidin (positive
control, 1.25 µM, 1× MIC). Data is the average of 2 biological × 2 techni-
cal repeats and error bars represent standard deviations. (b) Brightfield
and fluorescence imaging of B. cereus incubated for 15 minutes with 4%
DMSO (blank) or 1a (250 µM, 10× MIC). Absence of fluorescence indi-
cates that the cells are depolarized. Scale bars represent 10 µm.
bacteriostatic.61,62 The minimum bactericidal concentration
(MBC), defined as the lowest concentration needed to kill
99.9% of bacteria, was 35–40 µM for 1a, which is only slightly
higher than its MIC value (25–30 µM). This suggests that
compound 1a has bactericidal activity and points towards a
mode of action that involves the bacterial membrane. Further
evidence of a membrane-based mechanism came from live
cell imaging and Gram staining of the B. cereus bacteria after
24 h incubation with compound 1a (ESI†). This revealed a
pronounced elongation of the bacterial cells, which is a mor-
phological change that has been observed for other Bacillus
species upon alteration of their membrane composition.63 In
addition, we investigated the ability of 1a to cause membrane
depolarization of B. cereus cells using the voltage-sensitive
dye Disc3(5).64 This cationic membrane-permeable fluoro-
phore accumulates in polarized cells, where it self-quenches.
results confirm that 1a functions as a bactericidal agent
against B. cereus due to its ability to interact with the bac-
terial membrane.
Conclusions
When the membrane potential is dissipated, the dye is In this manuscript, we have identified a new crown ether urea
released into the medium and de-quenched, which can be derivative 1a that is able to selectively bind to the bacterial
followed by a fluorometric assay (Fig. 3a). Alternatively, the lipid PE over the mammalian lipid PC in both solution and in
depolarization event can also be studied using fluorescence liposomes. Furthermore, the compound functions as a bacteri-
imaging (Fig. 3b). In this case, cells that are polarized show a cidal agent against B. cereus with an MIC value of 25–30 µM
pronounced red fluorescence due to the accumulation of and causes membrane depolarization in this bacterium. The
Disc3(5), whereas depolarized cells do not show fluorescence. other urea and crown ether compounds in this manuscript did
Gramicidin was used as a positive control known to cause not have the same affinity for PE, demonstrating that the
membrane depolarization,65 and clindamycin was used as a 18-crown-6 and urea group are both required for strong PE
negative control because it targets the ribosome rather than head group binding and a rigid linker is needed between the
the bacterial membrane.66 At 25 µM (1× MIC) 1a caused two to achieve the right conformation. We are currently synthe-
partial depolarization of B. cereus, while full depolarization sizing a series of analogs of this promising lead compound to
was seen at 40 µM (1.6× MIC) and 250 µM (10× MIC). These optimize its antibacterial activity.
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19 M. M. Hashemi, B. S. Holden and P. B. Savage, in Fighting
Antimicrobial Resistance, Springer, Berlin, 2018, ch. 7, pp.
139–169.
Conflicts of interest
There are no conflicts to declare.
20 X.-Z. Lai, Y. Feng, J. Pollard, J. N. Chin, M. J. Rybak,
R. Bucki, R. F. Epand, R. M. Epand and P. B. Savage, Acc.
Chem. Res., 2008, 41, 1233–1240.
21 R. Joseph, A. Naugolny, M. Feldman, I. M. Herzog,
M. Fridman and Y. Cohen, J. Am. Chem. Soc., 2016, 138,
754–757.
Acknowledgements
SRH, EBJ, HG and NB would like to thank Tulane University
for start-up funds and the Louisiana Board of Regents for
funding (Grant No. LEQSF(2018-21)-RD-A-15). TD would like to 22 J. Choi, J. Kim, K. Kim, S.-T. Yang, J.-I. Kim and S. Jon,
thank the NSF-funded SMART-REU summer program
(NSF-DMR-REU Grant No. 1460637 and 1852275).
Chem. Commun., 2007, 1151–1153.
23 S. Yan, S. Chen, X. Gou, J. Yang, J. An, X. Jin, Y.-W. Yang,
L. Chen and H. Gao, Adv. Funct. Mater., 2019, 29, 1904683.
24 S. Guo, Q. Huang, Y. Chen, J. Wei, J. Zheng, L. Wang,
Y. Wang and R. Wang, Angew. Chem., 2021, 60, 618–
623.
Notes and references
1 K. Gould, J. Antimicrob. Chemother., 2016, 71, 572–575.
2 J. G. Hurdle, A. J. O’Neill, I. Chopra and R. E. Lee, Nat. Rev.
Microbiol., 2011, 9, 62–75.
25 C. L. Randle, P. W. Albro and J. C. Dittmer, Biochim.
Biophys. Acta, Lipids Lipid Metab., 1969, 187, 214–220.
26 M. P. Bos and J. Tommassen, Curr. Opin. Microbiol., 2004,
7, 610–616.
3 M. Zasloff, Nature, 2002, 415, 389.
4 W. Stillwell, An introduction to biological membranes: compo- 27 S. Hasim, D. P. Allison, B. Mendez, A. T. Farmer,
sition, structure and function, Elsevier, 2016.
5 R. M. Epand and R. F. Epand, J. Pept. Sci., 2011, 17, 298–
305.
D. A. Pelletier, S. T. Retterer, S. R. Campagna,
T. B. Reynolds and M. J. Doktycz, Front. Microbiol., 2018, 9,
219.
6 A. K. Marr, W. J. Gooderham and R. E. W. Hancock, Curr. 28 M. Vestergaard, N. A. Berglund, P. C. Hsu, C. Song,
Opin. Pharmacol., 2006, 6, 468–472.
7 D. R. Rice, K. J. Clear and B. D. Smith, Chem. Commun.,
2016, 52, 8787–8801.
8 C. M. Cooley, K. S. Hettie, J. L. Klockow, S. Garrison and
T. E. Glass, Org. Biomol. Chem., 2013, 11, 7387–7392.
9 V. E. Zwicker, B. L. Oliveira, J. H. Yeo, S. T. Fraser,
H. Koldso, B. Schiott and M. S. P. Sansom, ACS Omega,
2019, 4, 18889–18899.
29 H. Kawai, F. Chaudhry, A. Shekhar, A. Petrov, T. Nakahara,
T. Tanimoto, D. Kim, J. Chen, D. Lebeche and
F. G. Blankenberg, JACC Cardiovasc. Imaging, 2018, 11,
1823–1833.
G. J. L. Bernardes, E. J. New and K. A. Jolliffe, Angew. 30 A. Bandyopadhyay, K. A. McCarthy, M. A. Kelly and J. Gao,
Chem., 2019, 131, 3119–3123. Nat. Commun., 2015, 6, 6561–6561.
10 B. I. Murcar-Evans, A. D. Cabral, K. Toutah, E. D. de 31 M. Kelly, S. Cambray, K. A. McCarthy, W. Wang,
Araujo, A. Lai, P. M. Macdonald, A. Berger-Becvar,
D. Kraskouskaya and P. T. Gunning, Analyst, 2017, 142,
4511–4521.
E. Geisinger, J. Ortiz-Marquez, T. van Opijnen and J. Gao,
ACS Infect. Dis., 2020, 6, 2410–2418.
32 K. A. McCarthy, M. A. Kelly, K. Li, S. Cambray,
A. S. Hosseini, T. van Opijnen and J. Gao, J. Am. Chem. Soc.,
2018, 140, 6137–6145.
11 C.-L. Do-Thanh, M. M. Rowland and M. D. Best,
Tetrahedron, 2011, 67, 3803–3808.
12 L. H. Mak, S. N. Georgiades, E. Rosivatz, G. F. Whyte, 33 C. Dias, J. P. Pais, R. Nunes, M. T. Blazquez-Sanchez,
M. Mirabelli, R. Vilar and R. Woscholski, ACS Chem. Biol.,
2011, 6, 1382–1390.
13 E. Brunetti, S. Moerkerke, J. Wouters, K. Bartik and
I. Jabin, Org. Biomol. Chem., 2016, 14, 10201–10207.
14 J. O. Magrans, A. R. Ortiz, M. A. Molins, P. H. P. Lebouille,
J. T. Marques, A. F. Almeida, P. Serra, N. M. Xavier,
D. Vila-Vicosa, M. Machuqueiro, A. S. Viana, A. Martins,
M. S. Santos, A. Pelerito, R. Dias, R. Tenreiro,
M. C. Oliveira, M. Contino, N. A. Colabufo, R. F. M. de
Almeida and A. P. Rauter, Nat. Commun., 2018, 9, 4857.
J. Sánchez-Quesada, P. Prados, M. Pons, F. Gago and J. de 34 G. Townshend, G. S. Thompson, L. J. White, J. R. Hiscock
Mendoza, Angew. Chem., Int. Ed. Engl., 1996, 35, 1712–
and J. L. Ortega-Roldan, Chem. Commun., 2020, 56, 4015–
1715.
4018.
15 R. D. Hubbard, S. R. Horner and B. L. Miller, J. Am. Chem. 35 C. Dias, A. Martins, A. Pelerito, M. C. Oliveira, M. Contino,
Soc., 2001, 123, 5810–5811.
16 A. Alliband, F. A. Meece, C. Jayasinghe and D. H. Burns,
J. Org. Chem., 2013, 78, 356–362.
17 K. M. Bucholtz, P. C. Gareiss, S. G. Tajc and B. L. Miller,
Org. Biomol. Chem., 2006, 4, 3973–3979.
N. A. Colabufo and A. P. Rauter, Eur. J. Org. Chem., 2019,
2224–2233.
36 S. N. Tyuleva, N. Allen, L. J. White, A. Pépés,
H. J. Shepherd, P. J. Saines, R. J. Ellaby, D. P. Mulvihill and
J. R. Hiscock, Chem. Commun., 2019, 55, 95–98.
18 R. F. Epand, P. B. Savage and R. M. Epand, Biochim. 37 N. Allen, L. J. White, J. E. Boles, G. T. Williams, D. F. Chu,
Biophys. Acta, Biomembr., 2007, 1768, 2500–2509.
R. J. Ellaby, H. J. Shepherd, K. K. L. Ng, L. R. Blackholly,
3842 | Org. Biomol. Chem., 2021, 19, 3838–3843
This journal is © The Royal Society of Chemistry 2021

View Article Online
Organic & Biomolecular Chemistry
Communication
B. Wilson, D. P. Mulvihill and J. R. Hiscock, 52 J. M. Boon, R. Shukla, B. D. Smith, G. Licini and
ChemMedChem, 2020, 15, 2193–2205. P. Scrimin, Chem. Commun., 2002, 260–261.
38 V. Amendola, L. Fabbrizzi and L. Mosca, Chem. Soc. Rev., 53 J. C. McIntyre and R. G. Sleight, Biochemistry, 1991, 30,
2010, 39, 3889–3915. 11819–11827.
39 A. Späth and B. König, Beilstein J. Org. Chem., 2010, 54 R. J. Melander, D. V. Zurawski and C. Melander,
6, 32. MedChemComm, 2018, 9, 12–21.
40 N. Busschaert, S. J. Bradberry, M. Wenzel, C. J. E. Haynes, 55 M. Nahaie, M. Goodfellow, D. Minnikin and V. Hajek,
J. R. Hiscock, I. L. Kirby, L. E. Karagiannidis, S. J. Moore, Microbiology, 1984, 130, 2427–2437.
N. J. Wells, J. Herniman, G. J. Langley, P. N. Horton, 56 P. Bernat, K. Paraszkiewicz, P. Siewiera, M. Moryl, G. Płaza
M. E. Light, I. Marques, P. J. Costa, V. Félix, J. G. Frey and
P. A. Gale, Chem. Sci., 2013, 4, 3036–3045.
and J. Chojniak, World J. Microbiol. Biotechnol., 2016, 32,
157–157.
41 M. Kralj, L. Tušek-Božić and L. Frkanec, ChemMedChem, 57 M. A. Haque and N. J. Russell, Microbiology, 2004, 150,
2008, 3, 1478–1492. 1397–1404.
42 D. B. Hibbert and P. Thordarson, Chem. Commun., 2016, 58 N. A. Logan, J. Appl. Microbiol., 2012, 112, 417–429.
52, 12792–12805.
43 J. M. Seddon, Biochim. Biophys. Acta, Rev. Biomembr., 1990,
1031, 1–69.
59 E. Helgason, O. A. Økstad, D. A. Caugant, H. A. Johansen,
A. Fouet, M. Mock, I. Hegna and A.-B. Kolstø, Appl. Environ.
Microbiol., 2000, 66, 2627.
44 A. Chattopadhyay, Chem. Phys. Lipids, 1990, 53, 1–15.
45 D. Hoekstra, Biochemistry, 1982, 21, 1055–1061.
46 S. Das and P. Purkayastha, Langmuir, 2017, 33, 7281–7287.
47 A. J. M. Driessen, H. W. van den Hooven, W. Kuiper,
60 M. Balouiri, M. Sadiki and S. K. Ibnsouda, J. Pharm. Anal.,
2016, 6, 71–79.
61 F. Baquero and B. R. Levin, Nat. Rev. Microbiol., 2020, 19,
123–132.
M. Van de Camp, H.-G. Sahl, R. N. H. Konings and 62 M. Mahlapuu, J. Håkansson, L. Ringstad and C. Björn,
W. N. Konings, Biochemistry, 1995, 34, 1606–1614. Front. Cell. Infect. Microbiol., 2016, 6, 194.
48 T. O. Khatib, H. Stevenson, M. R. Yeaman, A. S. Bayer and 63 L. I. Salzberg and J. D. Helmann, J. Bacteriol., 2008, 190,
A. Pokorny, Antimicrob. Agents Chemother., 2016, 60, 5051. 7797.
49 K. Matsuzaki, O. Murase, N. Fujii and K. Miyajima, 64 J. D. te Winkel, D. A. Gray, K. H. Seistrup, L. W. Hamoen
Biochemistry, 1996, 35, 11361–11368. and H. Strahl, Front. Cell Dev. Biol., 2016, 4, 29.
50 J. M. Boon and B. D. Smith, J. Am. Chem. Soc., 1999, 121, 65 D. A. Kelkar and A. Chattopadhyay, Biochim. Biophys. Acta,
11924–11925. Biomembr., 2007, 1768, 2011–2025.
51 H. M. McConnell and R. D. Kornberg, Biochemistry, 1971, 66 J. Spížek and T. Řezanka, Appl. Microbiol. Biotechnol., 2004,
10, 1111–1120.
64, 455–464.
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