A selective membrane-targeting repurposed antibiotic with activity against persistent methicillin-resistant Staphylococcus aureus

Article abstract of DOI:10.1073/pnas.1904700116

Treatment of Staphylococcus aureus infections is complicated by the development of antibiotic tolerance, a consequence of the ability of S. aureus to enter into a nongrowing, dormant state in which the organisms are referred to as persisters. We report that the clinically approved anthelmintic agent bithionol kills methicillinresistant S. aureus (MRSA) persister cells, which correlates with its ability to disrupt the integrity of Gram-positive bacterial membranes. Critically, bithionol exhibits significant selectivity for bacterial compared with mammalian cell membranes. All-atom molecular dynamics (MD) simulations demonstrate that the selectivity of bithionol for bacterial membranes correlates with its ability to penetrate and embed in bacterial-mimic lipid bilayers, but not in cholesterol-rich mammalian-mimic lipid bilayers. In addition to causing rapid membrane permeabilization, the insertion of bithionol increases membrane fluidity. By using bithionol and nTZDpa (another membraneactive antimicrobial agent), as well as analogs of these compounds, we show that the activity of membrane-active compounds against MRSA persisters positively correlates with their ability to increase membrane fluidity, thereby establishing an accurate biophysical indicator for estimating antipersister potency. Finally, we demonstrate that, in combination with gentamicin, bithionol effectively reduces bacterial burdens in a mouse model of chronic deep-seated MRSA infection. This work highlights the potential repurposing of bithionol as an antipersister therapeutic agent.

Full text of DOI:10.1073/pnas.1904700116

A selective membrane-targeting repurposed antibiotic
with activity against persistent methicillin-resistant
Staphylococcus aureus
Wooseong Kima, Guijin Zoub, Taylor P. A. Haric, Ingrid K. Wiltc, Wenpeng Zhub, Nicolas Galleb, Hammad A. Faizid,
Gabriel L. Hendricksa, Katerina Toria, Wen Pana, Xiaowen Huanga,e, Andrew D. Steelec, Erika E. Csataryc,
Madeline M. Dekarskec, Jake L. Rosenc, Noelly de Queiroz Ribeirof, Kiho Leea, Jenna Porta, Beth Burgwyn Fuchsa,
Petia M. Vlahovskag, William M. Wuestc, Huajian Gaob, Frederick M. Ausubelh,i,1, and Eleftherios Mylonakisa,1
aDivision of Infectious Diseases, Rhode Island Hospital, Warren Alpert Medical School of Brown University, Providence, RI 02903; bSchool of Engineering,
Brown University, Providence, RI 02903; cDepartment of Chemistry and Emory Antibiotic Resistance Center, Emory University, Atlanta, GA 30322;
dDepartment of Mechanical Engineering, Northwestern University, Evanston, IL 60208; eDepartment of Dermatology, Nanfang Hospital, Southern Medical
University, Guangzhou 510515, China; fDepartment of Microbiology, Instituto de Ciências Biológicas, Universidade Federal de Minas Gerais, Belo Horizonte,
MG 31270-901, Brazil; gDepartment of Engineering Sciences and Applied Mathematics, Northwestern University, Evanston, IL 60208; hDepartment of
i
Molecular Biology, Massachusetts General Hospital, Boston, MA 02114; and Department of Genetics, Harvard Medical School, Boston, MA 02115
Contributed by Frederick M. Ausubel, June 17, 2019 (sent for review March 22, 2019; reviewed by Dale L. Boger and Ruhong Zhou)
Treatment of Staphylococcus aureus infections is complicated by
the development of antibiotic tolerance, a consequence of the
ability of S. aureus to enter into a nongrowing, dormant state in
which the organisms are referred to as persisters. We report that
the clinically approved anthelmintic agent bithionol kills methicillin-
resistant S. aureus (MRSA) persister cells, which correlates with its
ability to disrupt the integrity of Gram-positive bacterial membranes.
Critically, bithionol exhibits significant selectivity for bacterial com-
pared with mammalian cell membranes. All-atom molecular dynam-
ics (MD) simulations demonstrate that the selectivity of bithionol for
bacterial membranes correlates with its ability to penetrate and em-
bed in bacterial-mimic lipid bilayers, but not in cholesterol-rich
mammalian-mimic lipid bilayers. In addition to causing rapid mem-
brane permeabilization, the insertion of bithionol increases mem-
brane fluidity. By using bithionol and nTZDpa (another membrane-
active antimicrobial agent), as well as analogs of these compounds,
we show that the activity of membrane-active compounds against
MRSA persisters positively correlates with their ability to increase
membrane fluidity, thereby establishing an accurate biophysical in-
dicator for estimating antipersister potency. Finally, we demonstrate
that, in combination with gentamicin, bithionol effectively reduces
bacterial burdens in a mouse model of chronic deep-seated MRSA
infection. This work highlights the potential repurposing of bithionol
as an antipersister therapeutic agent.
has sparked new interest in membrane-active antimicrobial
therapeutic agents (7). The lipophilic tail of daptomycin, a nat-
ural cyclic lipopeptide synthesized by Streptomyces roseosporus,
inserts into Gram-positive bacterial membranes, forming oligo-
meric pores, causing membrane depolarization, potassium ion
efflux, and rapid cell death (8). Despite strong antimicrobial
potency against growing bacterial cells, daptomycin has not been
reported to be effective against persisters (9, 10).
Recently, our laboratory described the identification of membrane-
active synthetic retinoids that are efficacious in killing MRSA
Significance
There is a critical lack of therapeutic agents to treat infections
caused by nongrowing persister forms of methicillin-resistant
Staphylococcus aureus (MRSA). Although membrane-disrupting
agents can kill persister cells, their therapeutic potential has
been mostly overlooked because of low selectivity for bacterial
versus mammalian membranes. We report that the clinically
approved anthelmintic drug bithionol kills MRSA persisters by
disrupting membrane lipid bilayers at concentrations that ex-
hibit low levels of toxicity to mammalian cells. The selectivity
of bithionol results from the presence of cholesterol in mam-
malian but not in bacterial membranes. We also show that the
antipersister potency of membrane-active antimicrobial agents
correlates with their ability to increase membrane fluidity. Our
results significantly enhance our understanding of bacterial
membrane disruption and membrane selectivity.
MRSA bacterial persister drug repurposing membrane-active
|
|
|
antimicrobials membrane selectivity
|
taphylococcus aureus is a Gram-positive opportunistic human
S
pathogen carried by approximately one third of the human
Author contributions: W.K., G.Z., T.P.A.H., I.K.W., W.Z., N.G., H.A.F., G.L.H., W.M.W., H.G.,
F.M.A., and E.M. designed research; W.K., G.Z., T.P.A.H., I.K.W., W.Z., N.G., H.A.F., G.L.H.,
K.T., W.P., X.H., N.d.Q.R., K.L., J.P., and B.B.F. performed research; T.P.A.H., I.K.W., A.D.S.,
E.E.C., M.M.D., J.L.R., B.B.F., P.M.V., W.M.W., H.G., F.M.A., and E.M. contributed new
reagents/analytic tools; W.K., G.Z., T.P.A.H., I.K.W., W.Z., N.G., H.A.F., G.L.H., P.M.V.,
W.M.W., H.G., F.M.A., and E.M. analyzed data; and W.K., G.Z., W.M.W., H.G., F.M.A.,
and E.M. wrote the paper.
population. Despite antibiotic availability, S. aureus infections
are often hard to cure and remain one of the major causes of
death (1), in part because of the ability of S. aureus to enter into
a nongrowing antibiotic-tolerant state, in which the organisms
are referred to as persisters (2). Persisters exhibit significantly
reduced biosynthetic processes, which are the major targets for
most antibiotics (2). Persisters also exist in a metabolically low-
energy state (3) that prevents the energy-dependent uptake of
antibiotics such as aminoglycosides (4). S. aureus persisters are
present in high numbers in stationary-phase suspension cultures
and biofilms (4–6) and are responsible for chronic and relapsing
infections such as endocarditis, osteomyelitis, and prosthetic
implant infections (3).
Reviewers: D.L.B., The Scripps Research Institute; and R.Z., Columbia University.
Conflict of interest statement: F.M.A. and E.M. have financial interests in Genma Biosci-
ences and Octagon Therapeutics, companies that are engaged in developing antimicro-
bial compounds. E.M.’s and F.M.A.’s interests were reviewed and are managed by Rhode
Island Hospital (E.M.) and Massachusetts General Hospital and Partners HealthCare
(F.M.A.) in accordance with their conflict of interest policies. The remaining authors de-
clare no competing financial interests.
Published under the PNAS license.
1To whom correspondence may be addressed. Email: ausubel@molbio.mgh.harvard.edu
or emylonakis@lifespan.org.
Bacterial membranes are attractive antipersister targets be-
cause they can be disrupted independently of growth. Although
membrane-active agents are typically toxic to mammals because
of low membrane selectivity (7), the clinical success of daptomycin
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.
1073/pnas.1904700116/-/DCSupplemental.
www.pnas.org/cgi/doi/10.1073/pnas.1904700116
PNAS Latest Articles
|
1 of 6

persisters (6). They are also relatively nontoxic because they ex-
hibit a significant amount of selectivity for Gram-positive bacterial
membranes compared with mammalian membranes (6). Like
daptomycin, the antimicrobial retinoids also appear to insert into
Gram-positive bacterial membranes, causing rapid permeabilization
and cell death. In contrast to daptomycin, however, a subset of
the membrane-active synthetic retinoids also kill nongrowing
MRSA persister cells (6). In addition to the synthetic retinoids,
we have described 2 additional membrane-active compounds with
anti-MRSA activity: NH125, a histidine-kinase inhibitor (11), and
nTZDpa, a nonthiazolidinedione peroxisome proliferator-activated
receptor gamma partial agonist (12). Although NH125 and nTZDpa
have excellent anti-MRSA persister activity, they both cause
substantial hemolysis of red blood cells at high concentrations
(>32 μg/mL) (12, 13), which is a drawback for further devel-
opment as anti-MRSA therapeutic agents.
The membrane-active retinoids, NH125, and nTZDpa were
identified by screening ∼82,000 synthetic compounds by using a
high-throughput Caenorhabditis elegans–MRSA infection screen-
ing assay for compounds that block the ability of MRSA to kill the
nematodes (6, 11, 12). We subsequently screened 185 “hit” com-
pounds for the additional ability to permeabilize MRSA persisters
by using a SYTOX Green membrane permeability assay (11).
Another putative membrane-active antimicrobial identified in this
screen was the clinically approved anthelminthic drug bithionol,
which was chosen for additional studies because of its well-
established pharmacokinetic, toxicity, and safety profiles (14). In
this paper, we describe the therapeutic potential of bithionol as an
anti-MRSA persister antibiotic agent and provide a putative mode
of action that explains the ability of bithionol to selectively disrupt
bacterial compared with mammalian membrane lipid bilayers.
Moreover, by studying a panel of bithionol and nTZDpa analogs,
we found a strong correlation between the potency of anti-MRSA
persister activity and the ability to increase membrane fluidity.
A
B
Control
Bithionol
OH
Cl
OH
Cl
Cl
S
Cl
S. aureus MW2 persisters
C
DS. aureus MW2 biofilms
9
8
7
6
5
4
3
9
8
7
6
5
4
3
8
7
6
5
4
3
0
1
2
3
4
0
1
2
3
4
Time (h)
Time (h)
32 μg/mL BT 0 μg/mL BT
100 μg/mL Gm
200 μg/mL Lin
Non-treated
100 μg/mL Van
50 μg/mL Cipro
100 μg/mL Dap
16 μg/mL BT
8 μg/mL BT
S. aureus VRS1 Persisters
E 9
8
7
6
5
4
3
9
8
7
6
5
4
3
8
7
6
5
4
3
0
1
2
3
4
0
1
2
3
4
0
8
16
32
Time (h)
Time (h)
BT (μg/mL)
Non-treated
200 μg/mL Lin
100 μg/mL Dap
32 μg/mL BT
8 μg/mL BT
16 μg/mL BT
0 μg/mL BT
Fig. 1. Bithionol exhibits bactericidal activity against S. aureus persisters. (A)
Chemical structures of bithionol. (B) TEM micrographs showing the formation
of intracellular mesosome-like structures (red arrows), an abnormal cell di-
vision (brown arrow), or cell lysis (blue arrow) in S. aureus MW2 treated with
10× MIC (10 μg/mL) bithionol or 0.1% DMSO (control) for 2 h. (Scale bars,
500 nm.) (C and D) Viability of MRSA MW2 stationary-phase (C) or biofilm (D)
persister cells treated with 100× MIC of the conventional antibiotics vanco-
mycin (Van), gentamicin (Gm), ciprofloxacin (Cipro), daptomycin (Dap), line-
zolid (Lin), or the indicated concentrations of bithionol (BT) for 4 h (C) and
24 h (D), respectively. (E) The viability of S. aureus VRS1 stationary-phase
persisters treated with 100× MIC (200 μg/mL) linezolid (Lin), 100× MIC (100 μg/mL)
daptomycin (dap), or the indicated concentrations of bithionol (BT) as a
function of time. The data points on the x-axis are below the level of de-
tection (2 × 102 CFU/mL, or 2 × 102 CFU per membrane). Individual data points
(n = 3 biologically independent samples) are shown; error bars represent
means SD.
Results
Bithionol Shows Bactericidal Activity Against both Antibiotic-Resistant
S. aureus and S. aureus Persister Cells. Bithionol (Fig. 1A) was pre-
viously described as an antimicrobial agent with a minimal inhibitory
concentration (MIC) of ∼8 to 15 μg/mL against Gram-positive bac-
teria, including S. aureus (15). We confirmed that bithionol exhibits
antimicrobial activity against a variety of Gram-positive pathogens,
including vancomycin-resistant S. aureus (VRSA) and vancomycin-
and daptomycin-resistant enterococcal strains. In our hands, however,
bithionol was more potent than previously reported, with MICs of
0.5 to 2 μg/mL, comparable to daptomycin (SI Appendix, Table
S1). We also confirmed that bithionol exhibits relatively weak
antimicrobial activity against Gram-negative pathogens (MICs of
16 to >64 μg/mL; SI Appendix, Table S1).
within 2 h and 24 h, respectively (Fig. 1 C and D). In contrast,
MW2 persisters displayed a high level of tolerance to 100× MIC
of several conventional antibiotics, including daptomycin and
linezolid, which were tested for activity against planktonic per-
sisters. Similarly, bithionol killed vancomycin-resistant S. aureus
(VRSA) strain VRS1 persisters, whereas linezolid and daptomycin
exhibited no and nominal activity, respectively (Fig. 1E).
In contrast to a previous study that described bithionol as
bacteriostatic (16), we found that bithionol eradicated ∼107 CFU/mL
exponential-phase S. aureus strain MW2 within 3 h at 10 μg/mL
(10× MIC; SI Appendix, Fig. S1A). The rate of killing was
comparable to daptomycin (at 10× MIC) and significantly faster
than the killing kinetics of vancomycin (at 10× MIC; SI Appendix,
Fig. S1A). Consistent with its bactericidal activity, 10 μg/mL
bithionol caused a time-dependent decrease in optical density of
S. aureus cells comparable to the antiseptic detergent benzyldi-
methylhexadecylammonium chloride (16-BAC; SI Appendix, Fig.
S1B), indicating that bithionol has bacteriolytic activity. Indeed,
transmission electron micrographs (TEMs) showed that 10× MIC
bithionol disrupts MRSA membranes, causing the intracellular
formation of mesosome-like structures, abnormal cell divisions, and
cell lysis (Fig. 1B), which we previously observed when S. aureus
cells were treated with membrane-active antimicrobials (12).
We also found that bithionol killed stationary-phase MRSA
MW2 planktonic and biofilm persisters in a dose-dependent
manner and completely eradicated them at 32 μg/mL (32× MIC)
Bithionol Interacts with and Disrupts the Bacterial Mimetic Lipid
Bilayer. We performed all-atom molecular dynamics (MD) simu-
lations of bithionol interacting with simulated bacterial mem-
branes to elucidate a potential mechanism of action. The topology
and parameters of bithionol for the GROMOS54a7 force field
(17) were generated by Automated Topology Builder (18). We
used the previously established model of 1,2-dioleoyl-sn-glycero-3-
phosphocholine (DOPC)/1,2-dioleoyl-sn-glycero-3-phospho-
(1′-rac-glycerol) (DOPG) at a 7:3 ratio (19) to simulate negatively
charged S. aureus membranes (SI Appendix, Materials and Meth-
ods). The MD modeling showed that bithionol is initially recruited
to the membrane surface by the binding of polar hydroxyl and
chlorine groups to hydrophilic lipid heads (Fig. 2A and Movie S1).
After several hundred nanoseconds of sustained attachment,
bithionol penetrates into the membrane interior, maximizing in-
teractions between nonpolar benzene rings and hydrophobic lipid
tails (Movie S1). After penetration, bithionol embeds in the outer
leaflet of the lipid bilayer (Fig. 2A and Movie S1). The insertion of
2 of 6
|
www.pnas.org/cgi/doi/10.1073/pnas.1904700116
Kim et al.

indicating that bithionol interacts with and disrupts a bacterial
mimetic lipid bilayer.
A
B
7DOPC/3DOPG
7POPC/3chol
30
15
0
7DOPC/3DOPG
7POPC/3cholesterol
0 ns 290 ns
0 ns
314 ns
Bithionol Induces Rapid Membrane Permeabilization and an Increase
in Membrane Fluidity. By using a SYTOX Green permeability assay,
we found that, in contrast to daptomycin, bithionol induced dose-
dependent membrane permeability in both exponential-phase
MRSA MW2 cells and stationary-phase MRSA MW2 persisters
(Fig. 2D and SI Appendix, Fig. S1C). The fluorescence intensity
peaked at 4 μg/mL and then decreased at higher concentrations of
bithionol (Fig. 2D), most likely as a consequence of nucleic acids
released form lysed cells (20), consistent with the observation that
10 μg/mL bithionol lyses MRSA persisters (Fig. 1B). The dose-
dependent effects of bithionol on membrane permeability corre-
lated with its dose-dependent killing kinetics (Figs. 1C and 2D and
SI Appendix, Fig. S1C), suggesting that the bactericidal activity of
bithionol results from its membrane-disrupting activity.
Insertion of particular membrane disrupting compounds into
lipid bilayers is known to cause dramatic changes in membrane
fluidity that disrupts the normal liquid-crystalline phase of the
membrane (21–23). This results in passive permeabilization, loss
of membrane protein functions, leakage of cellular components,
and bacterial death (24). We therefore tested whether bithionol
alters MRSA MW2 membrane fluidity by utilizing the membrane
fluidity-sensitive dye Laurdan (25), which exhibits a fluorescence
emission wavelength shift dependent on the amount of adjacent
water molecules (25). Because water molecule penetration into
lipid bilayers is in turn determined by lipid packing density and
lipid bilayer fluidity, bacterial membrane fluidity can be quanti-
fied by using the Laurdan generalized polarization (GP) metric:
GP = (I440 − I490)/(I440 + I490), ranging from −1 (most fluid and
disordered) to +1 (most rigid and ordered) (25). As shown in
Fig. 2E, bithionol at concentrations greater than 8 μg/mL in-
duced a significant decrease in Laurdan GP in S. aureus MW2,
similar to 50 mM benzyl alcohol (BA), a known membrane flu-
idizer. These data are consistent with our observation that
bithionol concentrations greater than 16 μg/mL exhibited sig-
nificant killing activity against MRSA MW2 persisters (Fig. 1C).
117 ns
500 ns
168 ns
500 ns
-15
1
2
3
4
Distance from bilayer center
(nm)
C
D
Bithionol
DMSO
1 μg/ml 10 μg/ml 0.1%
S. aureus MW2
HKC-8
1000
800
600
400
200
0
1000
800
600
400
200
0
0
20
40
60
0
20
Time (min)
32μg/mL 16μg/mL 8 μg/mL 4 μg/mL
40
60
Time (min)
64 μg/mL
32 μg/mL
16 μg/mL
0 μg/mL
μg/mL
1 μg/mL
0 μg/mL
2
Bithionol
1 μg/ml 10 μg/ml 0.1%
DMSO
0.40
E
0.35
0.30
0.25
0.20
0.15
**
**
***
***
Fig. 2. Bithionol selectively disrupts bacterial lipid bilayers. (A) Representa-
tive configurations of MD simulations of bithionol from left to right: onset,
membrane attachment, membrane penetration, and equilibrium interacting
with 7DOPC/3DOPG or 7POPC/3cholesterol lipid bilayers. Bithionol and sodium
ions are depicted as large spheres, phospholipids are represented as chains,
and bonds in cholesterols are highlighted by thickened tubes. The atoms in
bithionol, phospholipids, cholesterol, and sodium ions are colored as follows:
hydrogen, white; oxygen, red; nitrogen, blue; sulfur, yellow; chlorine, green;
carbon, cyan; phosphorus, orange; and sodium, purple. For clarity, water
molecules are not shown. (B) The free-energy profiles of bithionol penetrating
into the indicated lipid bilayers as a function of the center-of-mass (COM)
distance to the bilayer. The dot-dashed blue and red lines mark the surface of
bacterial and mammalian membranes, respectively, averaged from the COM
locations of phosphate groups in the lipids of the outer leaflets. Error bars
represent means SD from 3 independent simulations. (C) GUVs consisting of
DOPC/DOPG (7:3) or POPC/cholesterol (7:3) labeled with 0.005% Liss Rhod PE
were treated with the indicated concentrations of bithionol or 0.1% DMSO
(control) and were monitored over time by using fluorescence microscopy.
(Scale bars, 10 μm.) (D) Uptake of SYTOX Green (Ex = 485 nm, Em = 525 nm)
by MRSA MW2 persister cells or human renal proximal cells (HKC-8) treated
with the indicated concentrations of bithionol. Results are shown as means;
n = 3 biologically independent samples. Error bars not shown for clarity. (E) S.
aureus MW2 membrane fluidity treated with the indicated concentrations of
bithionol for 1 h was evaluated based on Laurdan generalized polarization
(Laurdan GP). Laurdan GP = (I440 − I490)/(I440 + I490), where I440 and I490 are the
emission intensities at 440 and 490 nm, respectively, when excited at 350 nm.
The membrane fluidizer benzyl alcohol (50 mM) was used as a positive con-
trol. Individual data points (n = 3 biologically independent samples) are
Bithionol Does Not Penetrate Mammalian Membranes. We used MD
simulations to determine whether bithionol specifically penetrates
bacterial compared with mammalian membranes. Mammalian
membranes are comprised of phospholipids, sphingolipids, and
cholesterol (26), but can be modeled as a simplified bilayer
composed of the zwitterionic lipid 1,2-palmitoyl-oleoyl-sn-glycero-
3-phosphocholine (POPC) mixed with cholesterol ranging from 20
to 50 mol% (27, 28). At a POPC/cholesterol ratio of 7:3, MD
simulations using the GROMOS54a7 force field showed that
bithionol fails to penetrate the simulated mammalian bilayer (Fig.
2 A and B, SI Appendix, Fig. S2B, and Movie S2). Moreover, the
energy barrier and transfer energy for bithionol increases with
increasing percentages of cholesterol (SI Appendix, Fig. S3A).
These results are consistent with our observation that the presence
of cholesterol results in a more ordered alignment of the mem-
brane lipids (SI Appendix, Fig. S3D), as well as with configura-
tional and thermodynamic analyses (29, 30), which demonstrate
that cholesterol condenses the hydrophobic region of the mem-
brane (SI Appendix, Fig. S3B) and decreases membrane fluidity
(SI Appendix, Fig. S3C). An independent MD simulation using the
CHARMM force field (31) also showed that bithionol preferen-
tially penetrates bacterial-mimetic compared with mammalian-
mimetic lipid bilayers (SI Appendix, Fig. S4).
shown; error bars represent means
control and antibiotic treatment groups were analyzed by 1-way ANOVA and
post hoc Dunnett test (**P = 0.01 and ***P < 0.0001.).
SD. Statistical differences between
even a single bithionol molecule causes a substantial local increase
in lipid bilayer disorder (SI Appendix, Fig. S2 A and C). These MD
simulations demonstrate that the polarity of branch groups and
the hydrophobicity of core rings play important roles in membrane
attachment and penetration.
We further investigated the effects of bithionol on lipid bila-
yers by using biomembrane-mimicking giant unilamellar vesicles
(GUVs) consisting of a single lipid bilayer with a diameter of 10
to 100 μm. GUVs were constructed by using DOPC/DOPG lipids
at a 7:3 ratio as in the MD simulations. Lipid aggregates formed
on the surfaces of the GUVs exposed to 1 μg/mL bithionol, and,
Consistent with the MD simulations, bithionol did not cause
observable effects on GUVs formed with 7POPC/3cholesterol
(Fig. 2C and Movies S6–S8). Moreover, bithionol exhibited rel-
atively little hemolytic activity against human erythrocytes, with
an HC50 >64 μg/mL (SI Appendix, Fig. S5A), and did not induce
at 10 μg/mL, the GUVs burst (Fig. 2C and Movies S3–S5), SYTOX Green membrane permeabilization of HKC-8 human
Kim et al.
PNAS Latest Articles
|
3 of 6

renal proximal cells up to 64 μg/mL (Fig. 2D and SI Appendix,
Fig. S5 B and C).
the smaller chlorine and fluorine atoms, although the bromines may
be slightly disadvantageous for initial binding and penetration.
Bithionol in Combination with Gentamicin Shows Efficacy in a Mouse
Deep-Seated MRSA Infection Model. We previously showed that the
aminoglycoside antibiotic gentamicin, which is used to treat se-
vere chronic MRSA infections despite nephrotoxicity issues (33),
exhibits significant synergism with synthetic retinoids (6). Although
we found that relatively low concentrations of bithionol (8 μg/mL)
do not lead to a significant decrease in the viability of MRSA per-
sisters (Fig. 1 C and D), 8 μg/mL bithionol combined with as little as
2 μg/mL gentamicin completely eradicated MRSA MW2 stationary-
phase persister cells (Fig. 3A). Similarly, 8 μg/mL bithionol + 16 μg/mL
gentamicin eradicated biofilm persisters (Fig. 3B).
Structure–Activity Relationships. The MD simulations described
here earlier predicted that 2 key elements for the membrane ac-
tivity of bithionol are the initial binding to the membrane surface
using phenolic hydroxyl groups and membrane perturbation in-
duced by chlorinated benzene (Fig. 2A and Movie S1). To further
test this proposed mechanism and to obtain additional insights
into the effects of functional groups on the potency of bithionol,
we conducted structure–activity relationship (SAR) studies by
using a commercially available bithionol analog, Bitin-S, as well as
with 7 newly synthesized analogs (Table 1 and SI Appendix, Figs.
S6 and S7). The effect of binding affinity on antimicrobial and
membrane activity was tested by using Bitin-S, a sulfoxide de-
rivative of bithionol (32), and the bithionol methoxy analog BT-
OMe (Table 1). The polar sulfinyl group of Bitin-S provides ad-
ditional hydrophilic interactions with lipid heads. Bitin-S exhibited
decreased antimicrobial activity (MIC 8 μg/mL) and reduced
membrane activity (Table 1 and SI Appendix, Fig. S6). Reducing
polarity by substituting methoxy groups for the 2 hydroxyl groups
(BT-OMe; Table 1 and SI Appendix, Fig. S6) resulted in complete
We further tested the efficacy of bithionol in combination with
gentamicin in an MRSA mouse thigh infection model, which
mimics human deep-seated chronic infections (5). Consistent
with a previous study (5), vancomycin, gentamicin, or a combi-
nation of the 2 did not significantly reduce MRSA CFUs in the
mouse thigh (Fig. 3C), suggesting that the infecting bacterial
cells are persisters. Although 30 mg/kg bithionol alone showed
no effect on the viability of MRSA persisters, 30 mg/kg bithionol
combined with 30 mg/kg gentamicin killed ∼90% of the MRSA
nullification of both antimicrobial and membrane activity (Table 1 persister cells (P < 0.001; Fig. 3C). We evaluated the hepatic and
renal toxicity of bithionol in the mice in the experiments de-
scribed in Fig. 3C by measuring serum levels of alanine amino-
transferase (ALT) and blood urea nitrogen (BUN) (SI Appendix,
Fig. S8). Although the combination of vancomycin and genta-
micin significantly increase BUN levels (P < 0.01), the combi-
nation of bithionol and gentamicin increased neither.
and SI Appendix, Fig. S6), indicating that the phenolic hydroxyl
groups are critical for antimicrobial activity.
To test whether the size and polarization of the inserted mole-
cules can affect the extent of membrane perturbation (12), we
substituted alternative halogens for chlorine. Replacement of chlo-
rine with fluorine resulted in reduced membrane and antimicrobial
activities as seen with compounds BT-oF, BT-pF, and BT-opF, whereas
substitution with bromine showed similar membrane and antimi-
crobial activities as bithionol as demonstrated by BT-oBr, BT-pBr,
and BT-opBr (Table 1 and SI Appendix, Fig. S7). It is likely that the
polarity of the C-F bond in the fluoro derivatives may increase the
hydrophilicity of the aryl rings, thus decreasing membrane perme-
ability. The bromine derivatives showed increased energy barrier
values of 2 to 3 kBT (SI Appendix, Table S2), indicating that the
larger bromine atoms may cause more membrane perturbation than
The Killing of MRSA Persisters Is Correlated with Increased Membrane
Fluidity. The SAR studies reported here show that some com-
pounds that permeabilize MRSA persister cell membranes to
SYTOX Green, such as Bitin-S, BT-oF, or BT-pF, do not kill
them (Table 1 and SI Appendix, Figs. S6 and S7). We reported a
similar result in a previous SAR study on nTZDpa (12), where
we observed that nTZDpa-analogs 6 and 11 (Fig. 4 B and C)
induced rapid SYTOX Green permeabilization, but did not af-
fect the viability of MRSA persisters (12).
Interestingly, the bithionol SAR series showed that only
compounds leading to a significant increase in membrane fluidity
killed MRSA persisters (Fig. 4A, Table 1, and SI Appendix, Figs. S6
and S7). To test the hypothesis that there is a general correlation
between an increase in membrane fluidity and antipersister potency,
we measured the effect of nTZDpa and 11 nTZDpa derivatives
(Fig. 4 B and C) (12) on MRSA membrane fluidity. Consistent with
the bithionol analogs, only nTZDpa analogs that exhibit anti-
persister potency showed increased membrane fluidity at 32 μg/mL.
Furthermore, we observed substantial correlation between the
concentration of compound required for persister killing and the
amount of induced membrane fluidity as measured by Laurdan
GP for all 21 tested compounds at 32 μg/mL (R2 0.64, P of
slope < 0.0001; SI Appendix, Fig. S9). Consistent with these data,
a recent study reported that the antibiotic agent rhodomyrtone
causes increased membrane fluidity in Bacillus subtilis and kills
its persister cells (23).
Table 1. Structure–activity relationships for antibiotic activity
and membrane activity
R1
R3
R1
R3
OH
OH
R2
R2
Cl
S
Cl
R
Cl
Cl
bithionol
OH
R3
O
HO O HO
S
OH
OH
R3
AlCl3 (1 equiv)
DCM
22 ºC, 8 h
Zn dust (10 equiv)
AcOH
100 ºC, 4 h
R2
R2
R2
R2
R2
S
S
Cl Cl
+(1 equiv)
R3
R3
R3
Cmpd
R
R1
R2
R3
MIC
PKC
MP
MFI
Bithionol
S
OH
Cl
Cl
Cl
F
Cl
Cl
Cl
Cl
Cl
F
Br
F
Br
1
32
>64
>64
>64
32
>64
32
Yes
Yes
No
Yes
Bitin-S
SO
S
OH
OMe
OH
OH
OH
OH
OH
OH
8
No
BT-OMe
BT-oF
BT-oBr
BT-pF
BT-pBr
BT-opF
BT-opBr
>64
2
1
2
1
No
S
S
S
S
S
S
Yes
Yes
Yes
Yes
No
No
Yes
No
Yes
No
Yes
The data in this section show that the killing of MRSA per-
sisters is achieved only when membrane damage by membrane-
active agents is sufficiently severe to show increased membrane
fluidity, potentially making membrane fluidity a biophysical in-
dicator to identify and measure the potency of antipersister
antimicrobial agents.
Br
Cl
Cl
F
8
1
>64
32
Br
Yes
MIC, minimum inhibitory concentration (in micrograms per milliliter); PKC,
persister killing concentration (in micrograms per milliliter) required to kill
5 × 107 CFU/mL MRSA persister cells below the limit of detection; MP, memb-
rane permeabilization (determined based on SYTOX Green fluorescence in-
tensity); MFI, membrane fluidity increase (determined based on Laurdan GP
measurement).
Discussion
Membrane-active agents have attractive properties as antimi-
crobial agents, including fast killing rates, antipersister potency,
synergism with other antibiotic agents, and a low probability of
4 of 6
|
www.pnas.org/cgi/doi/10.1073/pnas.1904700116
Kim et al.

Appendix, Fig. S7). Replacement of the chlorines with bromines
decreased initial binding to the lipid bilayer (SI Appendix, Table
S2), but increased destabilization of the membrane compared with
chlorine and fluorine after penetration. Collectively, the antimicrobial
activity of bithionol can be modulated by binding affinity to lipid head
groups, penetration depth, and molecule size.
A
B
C
10
8
7
6
5
4
3
9
8
7
6
5
4
3
9
8
7
6
***
0
1
2
3
4
In contrast to bithionol and other membrane-active compounds
we have studied, daptomycin does not induce rapid SYTOX
Green membrane permeabilization or kill MRSA persisters
(Fig. 1C and SI Appendix, Fig. S1C). Importantly, we discovered
that bithionol, nTZDpa, and their analogs that exhibit anti-
MRSA persister potency induce SYTOX Green membrane per-
meabilization and an increase in membrane fluidity (Fig. 4, Table 1,
and SI Appendix, Figs. S6 and S7). These data are consistent with
previous reports that show that insertion of compounds into
membrane bilayers can increase membrane disorder and fluidity,
which subsequently causes passive membrane permeabilization (21,
22). Interestingly, we found that the initiation of SYTOX Green
membrane permeabilization occurs at a lower concentration of
the membrane-active compounds than is required to increase
membrane fluidity (Fig. 2 D and E and SI Appendix, Figs. S6 and
S7). Furthermore, some membrane-active agents, such as bitin-S,
bithionol fluorine analogs, and nTZDpa-analogs 6 and 11, that
induce SYTOX Green membrane permeabilization do not cause
a change in membrane fluidity and do not kill persisters (Fig. 4 B
and C and SI Appendix, Figs. S6 and S7). Bacterial membranes
consist of lipid rafts organized into microdomains having different
lipid compositions (35). Depending on lipid compositions, one
domain may be less ordered and more fluid, whereas another
domain may be more ordered and rigid (35). Because ordered and
rigid domains show more resistance to membrane active agents
(35) up to a certain threshold concentration, only less ordered
domains would be affected by membrane-active compounds, thus
making them SYTOX Green-permeable. However, this type of
Time (h)
8 μg/mL BT + 2 μg/mL Gm
8 μg/mL BT + 4 μg/mL Gm
8 μg/mL BT + 8 μg/mL Gm
8 μg/mL BT + 16 μg/mL Gm
8 μg/mL BT
16 μg/mL Gm
Non-treated
Fig. 3. Bithionol shows synergism with gentamicin against MRSA persisters
in vitro and in vivo. (A) Stationary-phase or (B) biofilm MRSA MW2 persisters
were treated with the indicated concentrations of bithionol (BT) combined
with gentamicin (Gm). Colony forming units (CFUs) were measured by serial
dilution and plating on TSA plates. The data points on the x-axis are below
the level of detection (2 × 102 CFU/mL, or 2 × 102 CFU per membrane). In-
dividual data points (n = 3 biologically independent samples) are shown;
error bars represent means SD. (C) Ten infected mice per group (n = 10
biologically independent animals) were treated with control (5% Killophor +
5% ethanol, i.p.), vancomycin (30 mg/kg, i.p.), gentamicin (30 mg/kg, s.c.),
bithionol (30 mg/kg, i.p.), or vancomycin (30 mg/kg, i.p.) or bithionol (30 mg/kg,
i.p.) combined with gentamicin (30 mg/kg, s.c.) every 12 h for 3 d at 24 h
postinfection. At 12 h after the last treatment, mice were euthanized.
Their thighs were excised and homogenized. CFUs from each mouse thigh
are plotted as individual points, and error bars represent the SD in each
experimental group. Statistical differences between control and antibiotic
treatment groups were analyzed by 1-way ANOVA and post hoc Tukey test
(***P < 0.001).
resistance selection (7). Unfortunately, most of these agents also
indiscriminately disrupt mammalian membranes. However, evolu-
tion has taken advantage of differences in bacterial and eukaryotic
membranes, as reflected in the production of cationic antimicro-
bial peptides by animals and plants that specifically target bacterial
cells. Gram-positive bacterial membrane lipid bilayers contain
∼25% anionic phospholipids, whereas mammalian membranes are
composed of zwitterionic (neutral) phospholipids and 20 to 50%
cholesterol (27, 34). Cationic antimicrobial peptides bind prefer-
entially to negatively charged bacterial membranes, and cholesterol
in animal membranes creates a condensing effect that confers
membrane rigidity and prevents the penetration of antimicrobial
peptides (34).
We found that bithionol penetration into negatively charged
bacterial mimetic lipid bilayers (7DOPC/3DOPG) is energeti-
cally favorable, whereas bithionol penetration into cholesterol-
rich mammalian mimetic lipid bilayers (7POPC/3cholesterol) is
energetically unfavorable (Fig. 2 A and B). Further, as the pro-
portion of cholesterol increases from 0 to 30%, the penetration
of bithionol becomes increasingly unfavorable (SI Appendix, Fig.
S3A), indicating that cholesterol plays a key role in bithionol’s
membrane selectivity.
MD modeling suggests that the 2 polarized phenolic hydroxyl
groups of bithionol play a major role in the presumed initial binding
to phospholipid headgroups via hydrogen bonding. SAR studies
showed that a methoxy analog of bithionol nullified bioactivity,
supporting this proposed mechanism of interaction (Table 1 and SI
Appendix, Table S2 and Fig. S6). In addition to providing presumed
polar interactions with lipid headgroups, the bithionol chlorine
moieties also appear to play a key role in lipid bilayer perturbation.
Briefly, after attachment, the binding affinity of bithionol is domi-
nated by hydrophobic interactions between its aromatic rings and
the hydrophobic tails of the membrane lipids, which drives the
penetration of the chlorinated benzene into the outer leaflet of the
lipid bilayer, causing lipid bilayer perturbation (SI Appendix, Fig. S2
A and C). Accordingly, the replacement of chlorine with fluorine
resulted in a decrease in antimicrobial potency (Table 1 and SI
A
C
4
R2
3
0.4
0.3
0.2
0.1
X
***
***
O
Cl
***
***
N
OH
2
3
4
R1
R1
Cmpd
R2
X
nTZDpa 4-Cl
H
S
S
S
>64
ns
B0.4
>64
ns
4
4-Cl
4-Cl
4-Cl
3,4-Cl
4-Cl
4-Cl
4-Cl
4-Cl
2,4-Cl
4-Cl
4-Cl
4-Cl
4-tBu
H
32
*
64
***
16
***
5
6
32
**
16
***
32
**
8
***
64
***
O
O
O
O
O
O
O
O
O
***
0.3
0.2
0.1
10
11
12
13
14
S21
S24
S26
H
8
***
4-Cl
3,4-Cl
4-Br
4-I
H
4-tBu
4-CF3
16
***
Fig. 4. Relationship between antipersister activity and alteration in mem-
brane fluidity. Membrane fluidity of (A) bithionol and its analogs or (B)
nTZDpa and its analogs at 32 μg/mL was evaluated by Laurdan GP. The
membrane fluidizer benzyl alcohol (B.A.; 50 mM) was used as positive con-
trol. (B) The blue-colored numbers above each bar indicate persister killing
concentration (PKC, in micrograms per milliliter) required to kill 5 × 107 CFU/mL
MRSA persister below the limit of detection (2 × 102 CFU/mL). (A and B)
Individual data points (n = 3 biologically independent experiments) are
shown; error bars represent means
SD. Statistical differences between
control and antibiotic treatment groups were analyzed by 1-way ANOVA
and post hoc Dunnett test; ns, no significance (P > 0.05), *P = 0.05, **P =
0.01, and ***P
< 0.001. Individual data points (n = 3 biologically in-
dependent experiments) are shown; error bars represent means
The structures of nTZDpa and its analogs.
SD. (C)
Kim et al.
PNAS Latest Articles
|
5 of 6

localized membrane damage may not be sufficient to cause an
overall increase in membrane fluidity. Over the threshold concen-
tration, most membrane domains would be disrupted, and, sub-
sequently, overall membrane fluidity would increase. It is also
possible that some compounds may attack only less ordered and
more fluid areas of membranes, which causes SYTOX Green
membrane permeability, but not an overall increase in membrane
fluidity. In any case, the killing of MRSA persisters is apparently
achieved only when the bacterial membrane is sufficiently dam-
aged to show increased membrane fluidity as detected by
Laurdan GP.
In conclusion, we report that the clinically approved anthel-
mintic bithionol is an effective antimicrobial agent against both
multidrug-resistant and -persistent Gram-positive pathogens.
Bithionol kills Gram-positive (but not Gram-negative) bacterial
cells by disrupting lipid bilayers while maintaining high selectivity
for bacterial compared with mammalian membranes, a conse-
quence of the presence of cholesterol in mammalian membranes.
Further, bithionol in combination with gentamicin effectively
eradicates S. aureus persisters in vitro and significantly reduces
bacterial burden in a mouse model of chronic deep-seated
MRSA infection. We also demonstrate that increased mem-
brane fluidity is a biophysical indicator to identify potent anti-
persister compounds. Our studies provide further understanding of
the molecular mechanisms by which membrane-active small
molecules selectively disrupt Gram-positive bacterial over
mammalian membranes and support the conclusion that
membrane-active antimicrobial agents have promising potential
to be used for treating chronic infections caused by bacterial
persisters.
Materials and Methods
SI Appendix, Materials and Methods describes in detail the materials and
procedures used in this study, including bacterial strains and growth con-
ditions, antimicrobial agents and chemicals, transmission electron micros-
copy, a minimal inhibitory concentration (MIC) assay, a killing kinetics assay,
a persister cell killing assay, a biofilm persister killing assay, a bacterial
membrane permeability assay, a mammalian membrane permeability assay,
a membrane fluidity assay, a human blood hemolysis assay, a giant uni-
lamellar vesicles (GUVs) assay, all-atom molecular dynamics (MD) simula-
tions, a deep-seated mouse thigh infection model, and general procedures
for the synthesis of bithionol analogs.
ACKNOWLEDGMENTS. This study was supported by National Institutes of
Health Grants P01 AI083214 (to F.M.A. and E.M.), P20 GM121344 (to
B.B.F.), and R35 GM119426 (to W.M.W.) and by National Science Foundation
Grant CMMI-1562904 (to H.G.). We thank the Institute of Chemistry and Cell
Biology (ICCB)–Longwood at Harvard Medical School for providing the
chemical libraries used in this study. We thank Dr. L. Rice for generously
providing the E. faecium strains. The simulations reported were performed
on resources provided by the Extreme Science and Engineering Discovery
Environment (XSEDE) through Grant MSS090046 and the Center for Compu-
tation and Visualization (CCV) at Brown University. The NMR instruments
used in this work were supported by National Science Foundation Grant
CHE-1531620.
1. A. S. Lee et al., Methicillin-resistant Staphylococcus aureus. Nat. Rev. Dis. Primers 4,
18033 (2018).
19. Y.-F. Chen, T.-L. Sun, Y. Sun, H. W. Huang, Interaction of daptomycin with lipid bi-
layers: A lipid extracting effect. Biochemistry 53, 5384–5392 (2014).
2. K. Lewis, Persister cells. Annu. Rev. Microbiol. 64, 357–372 (2010).
3. B. P. Conlon et al., Persister formation in Staphylococcus aureus is associated with ATP
depletion. Nat. Microbiol. 1, 16051 (2016).
4. K. R. Allison, M. P. Brynildsen, J. J. Collins, Metabolite-enabled eradication of bacterial
persisters by aminoglycosides. Nature 473, 216–220 (2011).
20. B. L. Roth, M. Poot, S. T. Yue, P. J. Millard, Bacterial viability and antibiotic susceptibility
testing with SYTOX green nucleic acid stain. Appl. Environ. Microbiol. 63, 2421–2431 (1997).
21. M. L. Cartron et al., Bactericidal activity of the human skin fatty acid cis-6-
hexadecanoic acid on Staphylococcus aureus. Antimicrob. Agents Chemother. 58,
3599–3609 (2014).
5. B. P. Conlon et al., Activated ClpP kills persisters and eradicates a chronic biofilm in-
fection. Nature 503, 365–370 (2013).
6. W. Kim et al., A new class of synthetic retinoid antibiotics effective against bacterial
persisters. Nature 556, 103–107 (2018).
7. J. G. Hurdle, A. J. O’Neill, I. Chopra, R. E. Lee, Targeting bacterial membrane function:
An underexploited mechanism for treating persistent infections. Nat. Rev. Microbiol.
9, 62–75 (2011).
8. W. R. Miller, A. S. Bayer, C. A. Arias, Mechanism of action and resistance to dapto-
mycin in Staphylococcus aureus and Enterococci. Cold Spring Harb. Perspect. Med. 6,
a026997 (2016).
9. B. Sharma, A. V. Brown, N. E. Matluck, L. T. Hu, K. Lewis, Borrelia burgdorferi, the
causative agent of lyme disease, forms drug-tolerant persister cells. Antimicrob.
Agents Chemother. 59, 4616–4624 (2015).
10. G. S. Sahukhal, S. Pandey, M. O. Elasri, msaABCR operon is involved in persister cell
formation in Staphylococcus aureus. BMC Microbiol. 17, 218 (2017).
11. W. Kim et al., Identification of an antimicrobial agent effective against methicillin-
22. Y. Wu et al., Antibacterial activity and membrane-disruptive mechanism of 3-p-trans-
coumaroyl-2-hydroxyquinic acid, a novel phenolic compound from pine needles of
Cedrus deodara, against Staphylococcus aureus. Molecules 21, E1084 (2016).
23. D. Saeloh et al., The novel antibiotic rhodomyrtone traps membrane proteins in
vesicles with increased fluidity. PLoS Pathog. 14, e1006876 (2018).
24. R. M. Epand, C. Walker, R. F. Epand, N. A. Magarvey, Molecular mechanisms of
membrane targeting antibiotics. Biochim. Biophys. Acta 1858, 980–987 (2016).
25. T. Parasassi, E. Gratton, Membrane lipid domains and dynamics as detected by
Laurdan fluorescence. J. Fluoresc. 5, 59–69 (1995).
26. T. Harayama, H. Riezman, Understanding the diversity of membrane lipid composi-
tion. Nat. Rev. Mol. Cell Biol. 19, 281–296 (2018).
27. M. Deleu, J.-M. Crowet, M. N. Nasir, L. Lins, Complementary biophysical tools to in-
vestigate lipid specificity in the interaction between bioactive molecules and the
plasma membrane: A review. Biochim. Biophys. Acta 1838, 3171–3190 (2014).
28. C. Hong, D. P. Tieleman, Y. Wang, Microsecond molecular dynamics simulations of
lipid mixing. Langmuir 30, 11993–12001 (2014).
resistant Staphylococcus aureus persisters using
strategy. PLoS One 10, e0127640 (2015).
a
fluorescence-based screening
29. F. de Meyer, B. Smit, Effect of cholesterol on the structure of a phospholipid bilayer.
Proc. Natl. Acad. Sci. U.S.A. 106, 3654–3658 (2009).
12. W. Kim et al., Discovery and optimization of nTZDpa as an antibiotic effective against
bacterial persisters. ACS Infect. Dis. 4, 1540–1545 (2018).
13. W. Kim et al., NH125 kills methicillin-resistant Staphylococcus aureus persisters by
lipid bilayer disruption. Future Med. Chem. 8, 257–269 (2016).
14. J. Keiser, J. Utzinger, Food-borne trematodiasis: Current chemotherapy and advances
with artemisinins and synthetic trioxolanes. Trends Parasitol. 23, 555–562 (2007).
15. F. S. Barr, G. F. Collins, L. G. Wyatt, Potentiation of the antimicrobial activity of bi-
thionol. J. Pharm. Sci. 54, 801–802 (1965).
16. A. L. Erlandson, Jr, C. A. Lawrence, Inactivating medium for hexachlorophene (G11) types
of compounds and some substituted phenolic disinfectants. Science 118, 274–276 (1953).
17. N. Schmid et al., Definition and testing of the GROMOS force-field versions 54A7 and
54B7. Eur. Biophys. J. 40, 843–856 (2011).
30. T. Róg, M. Pasenkiewicz-Gierula, I. Vattulainen, M. Karttunen, Ordering effects of
cholesterol and its analogues. Biochim. Biophys. Acta 1788, 97–121 (2009).
31. J. B. Klauda et al., Update of the CHARMM all-atom additive force field for lipids:
Validation on six lipid types. J. Phys. Chem. B 114, 7830–7843 (2010).
32. T. Meshi, M. Yoshikawa, Y. Sato, Metabolic fate of bis(3,5-dichloro-2-hydroxyphenyl)-
sulfoxide (bithionol sulfoxide). Biochem. Pharmacol. 19, 1351–1361 (1970).
33. C. Liu et al.; Infectious Diseases Society of America, Clinical practice guidelines by the
infectious diseases society of America for the treatment of methicillin-resistant
Staphylococcus aureus infections in adults and children. Clin. Infect. Dis. 52, e18–
e55 (2011).
34. J. R. Brender, A. J. McHenry, A. Ramamoorthy, Does cholesterol play a role in the
bacterial selectivity of antimicrobial peptides? Front. Immunol. 3, 195 (2012).
35. M. Bramkamp, D. López, Exploring the existence of lipid rafts in bacteria. Microbiol.
Mol. Biol. Rev. 79, 81–100 (2015).
18. A. K. Malde et al., An automated force field topology builder (ATB) and repository:
Version 1.0. J. Chem. Theory Comput. 7, 4026–4037 (2011).
6 of 6
|
www.pnas.org/cgi/doi/10.1073/pnas.1904700116
Kim et al.