Opposing Effects of PhoPQ and PmrAB on the Properties
of Salmonella enterica serovar Typhimurium:
Implications on Resistance to Antimicrobial Peptides
The increasing number of resistant bacteria is a major threat worldwide, leading to the search for new antibiotic agents. One of the leading strategies is the use of antimicrobial peptides (AMPs), cationic and hydrophobic innate immune defense peptides. A major target of AMPs is the bacterial membrane. Notably, accumulating data suggest that AMPs can activate the two-component systems (TCSs) of Gram-negative bacteria. These include PhoP-PhoQ (PhoPQ) and PmrA-PmrB (PmrAB), responsible for remodeling of the bacterial cell surface. To better understand this mechanism, we utilized bacteria deficient either in one system alone or in both and biophysical tools including fluorescence spectroscopy, single-cell atomic force microscopy, electron microscopy, and mass spectrometry (MoskowitzS. M.;Antimicrob. Agents Chemother.2012, 56, 1019−103022106224; ChengH. Y.;J. Biomed. Sci.2010, 17, 6020653976). Our data suggested that the two systems have opposing effects on the properties of Salmonella enterica. The knockout of PhoPQ made the bacteria more susceptible to AMPs by making the surface less rigid, more polarized, and permeable with a slightly more negatively charged cell wall. In addition, the periplasmic space is thinner. In contrast, the knockout of PmrAB did not affect its susceptibility, while it made the bacterial outer layer very rigid, less polarized, and less permeable than the other two mutants, with a negatively charged cell wall similar to the WT. Overall, the data suggest that the coexistence of systems with opposing effects on the biophysical properties of the bacteria contribute to their membrane flexibility, which, on the one hand, is important to accommodate changing environments and, on the other hand, may inhibit the development of meaningful resistance to AMPs.
Introduction<!>Materials and Methods<!>Peptide Synthesis and Purification<!>Fluorescent Labeling of
the Peptides<!>Construction of PmrAB Mutants<!>Antimicrobial Activity
of the Peptides<!>Extent of Membrane Polarization and Increase
in Membrane Permeability
of the Various Bacterial Strains<!>LPS Extraction
and Purification<!>Lipid-A Isolation and Matrix-Assisted Laser
Desorption/Ionization
Time-of-Flight (MALDI-TOF) Analysis<!>Peptide Binding to Whole
Bacteria<!>Atomic Force Microscopy Image Acquisition<!>AFM Image Analysis<!>Transmission
Electron Microscopy<!>Effect of PhoPQ and PmrAB
on the Susceptibility of the Bacteria
to AMPs<!><!>Bacterial Membrane Polarization<!><!>Permeability of the Bacteria
to SYTOX Green<!><!>Binding Affinity of Peptides<!><!>Transmission Electron Microscopy Images of the Bacteria<!><!>Transmission Electron Microscopy Images of the Bacteria<!>Extraction and Analysis of the Lipopolysaccharides<!><!>Cell Wall Properties Determined
by Single-Cell Atomic Force
Microscopy<!><!>Cell Wall Properties Determined
by Single-Cell Atomic Force
Microscopy<!><!>Cell Wall Properties Determined
by Single-Cell Atomic Force
Microscopy<!>Discussion<!>Effect of the Mutations
on the Extent of Bacterial Membrane
Polarization and Permeability<!>Effect of the Mutations on the Surface Charge<!>Bacterial Biophysical Properties:
Surface Density, Rigidity,
and Thickness of the Periplasmic Space<!>Statistical Analysis<!>Author Contributions<!>
<p>The battle between humans and pathogenic
microorganisms was thought1,2 to be developed once
antibiotics were discovered. Unfortunately,
the increasing number of resistant bacteria became a major threat
worldwide, leading to the search for new antibiotic agents.3−5 We now encounter bacteria resistant to multiple antibiotics with
different mechanisms of action.6−8 Therefore, there is an ongoing
effort to develop novel antibiotics with new mechanisms of action.
One of the leading strategies being investigated is the use of antimicrobial
peptides (AMPs), which are cationic and hydrophobic peptides used
by all types of living organisms as part of their innate immune system
targeting the bacterial surface.9−11</p><p>AMPs rapidly neutralize
a broad range of microbes, both Gram-negative
and Gram-positive bacteria, mainly by binding and perturbing their
cell envelope, which contains lipopolysaccharide (LPS) or lipoteichoic
acid (LTA), respectively, as well as their cytoplasmic membranes.12,13 Therefore, studying the remodeling mechanisms of the outer surface
is important to understand the interaction between bacteria and AMPs.14</p><p>Bacteria, being unicellular organisms,
possess several sensory
systems that are able to sense the environment and respond by activating
different genes. Two such systems in Gram-negative bacteria are the
two-component systems (TCSs) PhoP-PhoQ (PhoPQ) and PmrA-PmrB (PmrAB,
BasRS in Salmonella).15−17 In these systems, a
sensory protein/loop (PhoQ or PmrB, respectively), situated on the
inner membrane, is activated by an environmental signal, which in
turn phosphorylates a cytoplasmic protein (PhoP or PmrA, respectively)
that is able to induce several genes. These systems respond to changes
in the environmental concentrations of magnesium, iron, and aluminum
ions, pH,18−21 and AMPs.22−24 It is suggested that AMPs bind to the acidic Mg2+ domain on the membrane receptor PhoQ.25,26 In addition, histidine-rich AMPs were suggested to act as chelators
of magnesium ions, reducing Mg2+ concentration in the environment,
causing PhoPQ activation.23,27</p><p>In Salmonella enterica, PmrAB, on
the other hand, was not found to be directly activated by AMPs but
by the PhoPQ-activated gene, pmrD, which is known to directly activate
the PmrB response regulator, thereby activating PmrAB-regulated genes.28,29</p><p>Upon activation, PhoPQ and PmrAB induce many genes, some of
which
encode enzymes that modify the composition of the bacterial surface,
including LPS,30−33 in addition to other functions that are involved in pathogenicity.
For example, the enzymes encoded by pagP and pagL, which are induced
by PhoPQ, modify the lipid-A of the LPS, while the products of pmrE
and the pmrHJIFKLM operon, induced by PmrAB, modify its charge and
composition.34−36 Yet, many of the genes induced by these systems have
unknown functions.36 Because the purpose
of these systems is to sense and prepare the bacteria for the changing
environment, their combined action is believed to lead to the development
of resistance to AMPs.23,37</p><p>Indeed, the relevance of
these systems to the resistance of bacteria
to AMPs was demonstrated when phoP-KO or pmrA-KO bacteria displayed
modified susceptibility to some AMPs (particularly polymyxin B and
polymyxin E) compared to the wild-type (WT).38−40 However, despite
extensive studies, most of the work regarding these systems is of
a genetic nature.41−43</p><p>Surprisingly, although PmrAB can be activated
upon PhoPQ activation,
we have found that each of these systems has an opposite effect on
some aspects of bacterial biophysical properties and thus have a different
effect on resistance depending on the AMP. This might suggest that
each system confers resistance to a different kind of AMP, which may
have implications on the correlation between the mechanism of action
of AMP and how resistance is developed against it. Here, we investigated
the contribution of the PhoPQ and PmrAB systems to resistance to AMPs
in Salmonella typhimurium by utilizing
a biophysical approach to study bacterial rigidity, morphology, modification
of surface charge, membrane polarization, membrane permeation, and
cell wall density. By constructing bacteria lacking PhoP alone, PmrAB
alone, or both, we tried to shed light on the complex relationship
between these two systems. Surprisingly, although PmrAB can be activated
upon PhoPQ activation, we have found that each of these systems has
an opposite effect on some aspects of bacterial biophysical properties,
and each of the systems has a different effect on the resistance to
different AMPs, which might suggest that these systems confer resistance
to different kinds of AMPs, and it may imply to the mechanisms of
actions and resistance of different AMPs.</p><!><p>Rink amide 4-methylbenzhydrylamine (MBHA) resin and N-(9-fluorenyl)methoxycarbonyl (Fmoc)-amino acids were purchased from
Calbiochem-Novabiochem. Other reagents used for peptide synthesis
included trifluoroacetic acid (TFA, Sigma-Aldrich), N,N-diisopropylethylamine (DIEA, Sigma-Aldrich),
dichloromethane (DCM, peptide synthesis grade, Bio-Lab), dimethylformamide
(DMF, peptide synthesis grade, Bio-Lab), and hydroxybenzotriazole
(HOBT) and 2-(1H-benzotriazol-1-yl)-1,1,3,3-tetramethyluronium
hexafluorophosphate (HBTU) (peptide synthesis grade, Bio-Lab). Three S. enterica serovar Typhimurium strains ATCC 14028
(WT), the phoP-knockout derivative,44 a
gift from Prof. Shoshi Altuvia's lab (the Hebrew University,
Jerusalem), and a PmrAB knockout were used in the present study.
Kanamycin, tetracycline, and chloramphenicol were purchased from Sigma
(catalog no. P1004, K-1377, T-3383, C0378, respectively). The media
used were lysogeny broth (LB, containing 20 g/L LB broth, Conda),
SOC (26.6 g/L SOB, Conda, supplemented with 0.4% glucose, Merck),
and modified N-minimal media (MNMM, 5 mM KCl, 7.5 mM (NH4)2SO4, 0.5 mM K2SO4, 1 mM KH2PO4, 0.01 mM Tris–HCl pH 7.4, 1 mM MgCl2, 0.4% glucose, 38 mM glycerol, and 0.1% casamino acids).45</p><!><p>The peptides were
synthesized by a solid phase method on a rink amide MBHA resin using
an ABI 433A automatic peptide synthesizer (Applied Biosystems). The
resin-bound peptides (0.1 mequiv each) were cleaved from the resins
by trifluoroacetic acid (TFA), washed with dry ether, and extracted
with acetonitrile/water (50% v/v). All peptides were amidated at their
C terminus. The peptides were further purified by reverse-phase high-pressure
liquid chromatography (RP-HPLC) on a C18 reverse-phase
Bio-Rad column (250 × 10 mm2, 300 Å pore size,
5 μm particle size). A linear gradient of 10–90% acetonitrile
in water containing 0.1% TFA (v/v) in 40 min was used at a flow rate
of 1.8 mL/min. Each crude peptide contained one major peak, as revealed
by RP-HPLC. The purified peptides were shown to be homogeneous (>98%)
by analytical HPLC. Electrospray mass spectroscopy confirmed their
identity.23</p><!><p>Resin-bound peptides
with a free N-terminal amino group were treated with rhodamine B N-hydroxysuccinimide dissolved in dimethylformamide (DMF)
containing 3% diisopropyl ethyl amine for 24 h. The resin-bound rhodamine
was then washed thoroughly with DMF and then with DCM, dried, and
then cleaved using TFA. The peptides were purified (greater than 98%
homogeneity) by RP-HPLC on a C4 column using a linear gradient
of 10–90% acetonitrile in 0.1% TFA for 40 min. The peptides
were subjected to amino acid and mass spectrometry analysis to confirm
their composition.</p><!><p>Mutants
in the PmrAB
system were created using the method of red recombinase.1 In short, S. typhimurium WT were transformed with the pKD46 plasmid after being passed in
a S. enterica LB5010 restriction mutant.
The bacteria were then grown in LB with ampicillin at 28 °C.
Colonies were picked, grown in LB with ampicillin and 0.4% arabinose
at 28 °C and made competent for electroporation using multiple
washes in sterile distilled water with 10% glycerol and frozen at
−80 °C until use. The DNA for knockout of the bacteria
was composed of the gene for tetracycline together with its promoter,
taken from pRS551, with 50bp flanking regions taken from the 5′
and 3′ of pmrA and pmrB accordingly (in Salmonellae, these genes are termed basS and basR). This fragment was inserted
into a pSUPER plasmid, passed in a S. enterica LB5010 restriction mutant, and cut using restriction enzymes. The
bacteria were then electrically transformed with the linear DNA product.
The bacteria were grown for 2 h in SOC supplemented with 0.4% arabinose
at 37 °C and then plated on SOC plates with tetracycline and
grown at 37 °C. Colonies that grew on tetracycline but not on
ampicillin were assayed using polymerase chain reaction (PCR) with
one primer at the 5′ end of tetracycline and one at the 3′
end upstream of the inserted construct in the Salmonellae genome. PCRs of positive colonies were sent for sequencing to ensure
that the pmrAB genes were replaced with tetracycline. The PmrAB knockout
in the WT was termed PmrAB-KO, and the PmrAB knockout in the phoP-KO
background was termed double-KO (DKO).</p><!><p>The antimicrobial
activity of the peptides was examined in sterile 96-wells plates (Nunc
F96 microtiter plates) in a final volume of 100 Î¼L as follows.
Aliquots (50 Î¼L) of an overnight suspension-containing bacteria
(late-log phase diluted 1:5000) were added to 50 Î¼L of modified
N-minimal media (MNMM) containing the peptides in a serial twofold
dilution, and antibiotics, if required, to select for the specific
mutant. Inhibition of growth was determined by the visibility of turbidity
in comparison to blank with no growth after an incubation of 18–20
h at 37 °C with agitation. Antibacterial activity was expressed
as the minimal inhibitory concentration (MIC) in which no growth could
be observed.</p><!><p>Typhimurium ATCC 14028 WT
and the mutants were grown to the mid-log phase, centrifuged, and
suspended in sodium phosphate buffer (SPB) (8 mM Na2HPO4, 2 mM NaH2PO4, pH = 7.4). Their OD
was measured, and they were diluted to reach OD600nm =
0.1 in 10 mL. Two microliters of SYTOX Green (for membrane permeability,
Invitrogen, S7020) or 2 μL of 3,3′-dipropylthiadicarbocyanine-iodide
[DiSC2(5), for membrane potential, Molecular Probes (Junction
City, OR)] was added to the bacterial stock to a final concentration
of 1 Î¼M, and they were incubated with agitation for 10 min (membrane
permeability) or 20 min (membrane potential) at room temperature.
The bacteria were transferred to a black 96-wells plate (Nunc) (containing
an AMP at different concentrations for membrane penetration), and
a kinetic measurement was performed over 20 min using a BioTek multiplate
reader (Synergy HT) fluorescence detector (SYTOX Green: ex = 486 nm,
em = 528 nm; DisC2(5): ex = 620 nm, em = 680 nm).</p><!><p>The cells for LPS isolation
were grown in 1 L LB medium supplemented with 1 mM MgCl2 and the appropriate antibiotic at 37 °C until OD600nm = 0.8 and subsequently harvested (7000 rpm, 15 min, room temperature).
The pellet was resuspended in 10 mL of di-distilled water (DDW) and
lyophilized. LPS isolation was performed according to the protocol
of Darveau and Hancock.46 This method is
based on the mechanical disruption of the cells, followed by sodium
dodecyl sulfate (SDS) solubilization and magnesium precipitation of
the LPS in cold ethanol. For cell breakage, the Precellys tissue homogenizer
and the Precellys glass kit 0.1 mm (Peqlab Biotechnologie GmbH, Erlangen,
Germany) were used instead of a French pressure cell. The yield of
LPS obtained was quantified by the amount of 2-keto-3-deoxyoctonate
(KDO) recovered relative to the cell dry weight.47 KDO purchased from Sigma was used for a standard curve.
For visualization, 5 Î¼g of LPS was loaded on SDS-polyacrylamide
gel electrophoresis (PAGE) (15% resolving gel, 4% stacking gel) and
stained using the sensitive silver staining method of Tsai and Frasch.48</p><!><p>To detect modifications
in the lipid-A structure, lipid-A was isolated from LPS by mild hydrolysis,
as described previously.49 All matrix-assisted
laser desorption/ionization time-of-flight (MALDI-TOF) analyses were
carried out in the negative-ion mode on a Bruker Biflex III MALDI-TOF
(Bruker Daltonics, Bremen, Germany). Herein, we used the AB SCIEX
5800 MALDI-TOF/TOF System equipped with a Nd:YAG (355 nm) laser with
1 kHz pulse in a negative-ion mode to analyze the lipid-A composition
of the different strains. Briefly, 0.5–1 mg of purified LPS
was dissolved in 500 Î¼L of 1% SDS in 10 mM sodium acetate buffer
(pH 4.5) and incubated in an ultrasonic bath for 10 min. The samples
were hydrolyzed at 100 °C for 1 h and then dried in a SpeedVac
vacuum concentrator. The samples were washed with 100 Î¼L of
distilled water and 500 Î¼L of acidified ethanol (100 Î¼L
of 4 M HCl was mixed with 20 mL of 95% ethanol) and centrifuged (2000g, 10 min, room temperature). The pellets obtained were
washed again with 500 Î¼L of 95% ethanol. After repeating the
washing steps, lipid-A was finally resuspended in 100 Î¼L of
distilled water and lyophilized. As a matrix, we used 20 mg/mL 5-chloro-2-mercaptobenzothiazole
(CMBT, Sigma-Aldrich) dissolved in methanol–chloroform (1:1).
Typically, 300 shots were accumulated for each mass spectrum. The
spectra were calibrated externally using the quantified peptides used
as peptide standard (Bruker Daltonics).</p><!><p>The Salmonella WT and mutants were
grown to mid-log in LB with the appropriate
antibiotics, washed twice with phosphate-buffered saline (PBS), and
diluted to OD600nm = 1 in PBS. Rhodamine-labeled peptides
were prepared at 3 Î¼M, and 20 Î¼L was dispensed to each
tube. Each strain (180 Î¼L) was added to the tubes to complete
the volume to 200 Î¼L with the peptides at a final concentration
of 0.3 μM. The bacteria were incubated for 10–15 min
and centrifuged at 14 000 rpm for 5 min. Fifty microliters
from the supernatant was added to 50 μL of guanidinium–HCl
6 M (with 20 mM ethylenediaminetetraacetic acid (EDTA), 50 mM Tris–HCl,
pH 6.5), and the fluorescence was read using a BioTek multiplate reader
(Synergy HT) fluorescence detector (ex = 529 nm, em = 590 nm). The
percentage of binding was calculated using tubes with the peptides
but without bacteria for 0% binding. Treatments were done in triplicate.</p><!><p>AFM imaging
was performed as described previously.50 Prior to the measurements, bacterial culture was grown to the mid-log
phase, washed two times in PBS, resuspended in PBS, and adjusted to
OD600nm of 0.1. Bacteria were incubated for 4 h on freshly
cleaved mica sheets coated with poly-l-lysine (0.01 mg/mL,
P4832, Sigma) and fixated with 2% glutaraldehyde for 20 min. The samples
were washed again with filtered DDW to remove glutaraldehyde traces
and left to dry overnight at room temperature. Images of bacteria
were acquired with MultiMode AFM (Bruker, Santa Barbara, CA) equipped
with a NanoScope V controller and a small scanner. Images were recorded
in air, at room temperature (22–24 °C), in the PeakForce
quantitative nanomechanical mapping (QNM) mode using silicon nitride
WS-levers (ORC8-PS-W, Olympus) with a nominal spring constant of 0.76
N/m. The PeakForce QNM AFM imaging mode yields quantitative nanomechanical
mapping of material properties, including DMT modulus of elasticity
(rigidity). At the same time, sample topography was imaged with high
resolution (1024 pixels) and minimized sample distortion due to the
fine adjustment of the force applied to the sample surface. The applied
force was adjusted around 1 nN, and the scan rate was set to 1 Hz.
Imaging was carried out at different scales to verify the consistency
and robustness of the evaluated structures. Numerical data presented
are the mean value (±standard deviation (SD)) of DMT modulus
of four 150 × 150 nm2 samples, taken from a 700 ×
700 nm2 field (n ≥ 8 cells for
each treatment).</p><!><p>The PeakForce
QNM (Bruker, Santa
Barbara, CA) mode enables acquiring topography images with elasticity mapping simultaneously. Representative
images are presented in Figure 4. The topography of the bacterial cell surface is shown in Figure 4A. In addition, the
estimations of bacterial cell length were done as distances between
the left and right inflection points in the profiles taken, along
the cell (Figure 4C).
The elasticity of the bacterial cell wall was evaluated as the mean
over a certain area at the DMT modulus map (Figure 4B). The areas for analysis were chosen to
be the highest and most uniform areas possible (black rectangles).
This was done to avoid the influence of cell surface geometry (cell
surface curvature) on the measured values. All measurements were carried
out with the original "NanoScope Analysis" software
(Bruker, Santa Barbara, CA). Elements of flooding analysis (Figure 4D) were determined
using WSxM software.51</p><!><p>The cells were centrifuged,
and the pellet was loaded on aluminum disks with a depth of 100 Î¼m
(Engineering Office M. Wohlwend GmbH, Switzerland) and covered with
a flat disc. The sandwiched sample was frozen in an HPM010 high-pressure
freezing machine (Bal-Tec, Liechtenstein). The cells were subsequently
freeze-substituted in an AFS2 freeze substitution device (Leica Microsystems,
Austria) in anhydrous acetone containing 2% glutaraldehyde and 0.2%
tannic acid osmium tetroxide for 3 days at −90 °C and
then warmed up to −30 °C over 24 h. The samples were washed
three times with acetone, incubated for 1 h at room temperature with
2% osmium tetroxide, washed three times with acetone, and infiltrated
for 5–7 days at room temperature in a series of increasing
concentrations of Epon in acetone. After polymerization at 60 °C,
60–80 nm sections were stained with uranyl acetate and lead
citrate and examined in a Tecnai T12 electron microscope (FEI, Holland)
operating at 120 kV, utilizing a 2k × 2k ES500W Erlangshen CCD
camera (Gatan, U.K.).</p><!><p>We investigated both natural and de novo-designed AMPs (Table 1). The list includes (i) the human cathelicidin LL-37, (ii) a de novo-designed 15-mer AMP designated K6L9 and (iii) its d,l-amino acid-containing
analogue, d,l-K6L9, and (iv) d,l amino acid histidine derivative, d,l-H6L9, (v) a 12-mer d,l-K5L7 and (vi) its caprylic-acid-conjugated
analogue C8-d,l-K5L7, and
(vii) the frog antimicrobial peptide Temporin L.23 These peptides have different net positive charges under
the experimental conditions, as well as hydrophobicity (represented
by RP-HPLC retention times), as indicated in Table 1. The peptides were tested for their antibacterial
activity against WT S. enterica serovar
Typhimurium ATCC 14028 (S. typhimurium) and its mutants (Table 2). The minimal inhibitory concentration (MIC) was defined
using a broth microdilution antimicrobial assay. S.
typhimurium have been widely used for the exploration
of bacterial resistance acquired from these two TCSs.52,53 To determine whether a short-term induction would increase the resistance
of the bacteria toward the peptides, which does not affect bacterial
growth, we grew the bacteria in the presence of the peptides at declining
concentrations and measured absorbance (A600) for cell growth. As shown in Table 2, phoP-KO was more susceptible to all AMPs compared
to the WT and showed a similar susceptibility to the C8-K5L7 lipopeptide. Surprisingly, the pmrAB-KO gained resistance
to four out of the seven peptides compared to the WT. However, similar
to the phoP-KO, it showed a higher susceptibility to the histidine-substituted
AMP. Interestingly, the double knockout (DKO) bacteria showed a susceptibility
picture that combines the susceptibility picture of both single mutants.
These results are probably attributed to the overall difference between
the biophysical properties of the WT compared to the mutants and their
susceptibilities toward the peptides.</p><!><p>All of the peptides
are amidated
at their C-termini. Underlined and bold font amino acids are d-enantiomers.</p><p>Relative
hydrophobicity is reflected
by the percent of acetonitrile at the retention time.</p><p>WT: S. enterica serovar Typhimurium 14 028s wild-type
phoP-KO; S. enterica serovar Typhimurium
14 028s phoP
knockout pmrAB-KO; S. enterica serovar
Typhimurium 14 028s PmrAB knockout DKO; S. enterica serovar Typhimurium 14 028s phoP and PmrAB knockout.</p><!><p>We examined the effect
of the TCSs on the ability of the bacteria to maintain membrane potential
using the potential-sensitive dye DisC2(5), whose fluorescence
is quenched due to the oligomerization of the dye upon binding to
the membrane of electrically polarized cells. The data shown in Figure 1 reveal that both
WT and pmrAB-KO are more polarized compared to phoP-KO and DKO bacteria,
albeit not strongly. This suggests that the membranes of the WT and
pmrAB-KO are less permeable compared to the membranes of the other
two mutants. In agreement with these results, the antimicrobial activities
(Table 2) of most peptides
toward the WT and the pmrAB-KO mutant are similar but lower compared
to phoP-KO and DKO, both of which also share similar susceptibilities
to most of the peptides. These results also agree with the higher
rigidity of the WT and the pmrAB-KO compared to phoP-KO and DKO (see
the sections on AFM studies)</p><!><p>Transmembrane potential of the WT and mutant Salmonellae. The WT and mutant Salmonellae were grown to the
mid-log growth phase and then washed twice with SPB. Then, the bacteria
were diluted to OD600nm = 0.1, and DisC2(5)
dye was added (1 Î¼M final concentration). The bacteria were
incubated with the dye for 20 min, and then the fluorescence was measured
(ex = 620 nm, em = 680 nm). All data represent mean ± SD from
three independent experiments. Error bars represent standard error.
One-way analysis of variance was used to analyze the data. Results
showed a statistically significant difference (*P-value <0.005).</p><!><p>The fluorescent
dye SYTOX Green was used to observe whether the KO of the TCSs has
an effect on the permeability of the bacterial membrane. The fluorescence
of this dye increases upon binding to nucleic acids, thus indicating
DNA binding and cell penetration. The results shown in Figure 2 represent the percentage of
fluorescence compared to the WT, taken as 100%. The data reveal that
the fluorescence of the WT and the pmrAB-KO mutant is considerably
less than that of the phoP-KO and DKO.</p><!><p>Membrane penetration
assay of the WT and mutant Salmonellae. The WT and
mutant Salmonellae were grown to the
mid-log growth phase and then washed twice with SPB. The bacteria
were diluted to OD600nm = 0.1, and SYTOX Green dye was
added (1 Î¼M final concentration). The bacteria were incubated
with the dye for 10 min. Then, the bacteria were added to a black
opaque 96-wells plate containing C8-d,l-K5L7 at different concentrations. The fluorescence was measured
(ex = 485 nm, em = 529 nm) after 3 min of incubation. All data represent
mean ± SD from three independent experiments. Error bars represent
standard error. One-way analysis of variance was used to analyze the
data. Results showed a statistically significant difference (*P < 0.001).</p><!><p>The differences in the
activities of the peptides on the bacteria could also be attributed
to differences in their binding to the bacteria. This property is
also influenced by the charge of the bacterial surface. To test this
possibility, we synthesized and fluorescently labeled a highly positively
charged peptide, pentalysine, with the fluorescent probe rhodamine.
A d-amino acid was incorporated into the peptide to disrupt
its conformation and eliminate the possible contribution of the structure
to the binding process. The bacteria were sedimented by centrifugation,
and the supernatant was sampled to measure the remaining fluorescence.
The fluorescence of the supernatant was compared to that of a control
without bacteria to calculate the percentage of binding. The data
shown in Figure 3 reveal
a similar extent of binding to WT and the pmrAB-KO, which is less
than that of phoP-KO and DKO but are also similar to each other. Importantly,
similar to the membrane polarization assay, these results correlate
with the antimicrobial activity (Table 2) since most peptides have the same activity on the
WT and the pmrAB-KO mutant, which is lower compared to phoP-KO and
DKO that also share similar susceptibilities to most of the peptides.</p><!><p>Binding
of KKkKK-rhodamine to the bacteria. Bacteria were grown
to the mid-log growth phase and diluted to OD600nm = 1
in PBS (−/−). Then, the lysine-rhodamine peptide was
added and incubated with the bacteria for 10 min. The bacteria were
then centrifuged at 14 000 rpm, and the supernatants were transferred
to a black 96-wells plate. The fluorescence of the supernatants was
measured (ex = 529 nm, em = 590 nm) and compared to a control without
bacteria to calculate the percent of binding. Results are an average
of five independent experiments (N = 4 per experiment).
Error bars represent standard error. One-way analysis of variance
was used to analyze the data. Results showed a statistically significant
difference (P < 0.002).</p><!><p>Resistance to AMPs could result from a change in the structure of
the cell wall as suggested for Staphylococcus aureus.54 High-pressure freezing and free substitution
followed by transmission electron microscope (TEM) were therefore
used to visualize cross sections of the bacteria, allowing us to visualize
the (i) inner and outer membranes, (ii) periplasmic space, and (iii)
cell wall thickness and density. Figure 4 shows the representative
images of the WT and mutants. As expected, the cell wall of all of
the bacteria contains two lipid layers seen as white lines. However,
an intense electron opaque outer layer and a dense periplasmic layer
are observed only in pmrAB-KO (Figure 4), which might explain the high rigidity of the cell
wall of pmrAB-KO compared to the other strains (Figure 5).</p><!><p>Imaging of the bacteria using transmission electron
microscopy
(TEM). Bacteria were imaged using TEM, and representative images were
chosen. The two membranes of these Gram-negative bacteria can be seen
as white thin lines, a stronger (darker) staining of the bacteria
means a denser array of proteins. (A) WT bacteria revealed a single
and uniform electron density staining at the periplasmic space and
a thin layer of highly stained material over the outer membrane. (B)
phoP-KO strain exhibited light staining of the membranes and almost
absent surface elements. (C) pmrAB-KO strain displayed a layer structure
with three distinct zones in its periplasmic space and a thick dark
layer beyond the outer membrane. (D) DKO strain resembles that of
the WT and phoP-KO strains but with thicker and longer external polysaccharides
and surface elements attached to its surface.</p><p>Comparison
of the rigidities of WT and mutant Salmonellae. All
values of rigidities are normalized to the WT (100%). Each
bar represents the corresponding mean value over 10 cells. All data
represent mean ± SD from three independent experiments. Error
bars represent standard error. One-way analysis of variance was used
to analyze the data. Results showed a statistically significant difference
(*P-value <0.005).</p><!><p>The TEM pictures of the WT bacteria revealed a single and uniform
electron density staining at the periplasmic space and a thin layer
of highly stained material over the outer membrane, commonly attributed
to surface-anchored proteins (Figure 4A). The phoP-KO strain exhibited light staining of
the membranes and almost absent surface elements (Figure 4B). Remarkably, the pmrAB-KO
strain displayed a layer structure with three distinct zones in its
periplasmic space, and a thick dark layer beyond the outer membrane,
caused by the high anionic charge of the LPS (Figure 4C). Interestingly, the staining pattern of
the DKO strain resembles that of the WT and phoP-KO strains (Figure 4D) but with thicker
and longer external polysaccharides due to the longer LPS chains and
surface elements attached to its surface.</p><p>In addition, we found
a substantial difference between the average
thickness of the periplasmic thickness between WT (12.0 ± 1.0
nm) and that of phoP-KO (14.0 ± 1.5 nm) and to a much lesser
extent with that of pmrAB-KO (20.0 ± 1.0 nm) and DKO mutants
(20.0 ± 1.1 nm).</p><!><p>Following the TEM results that showed changes in the cell wall,
as
well as the notion that the lipid composition of the LPS can be affected
by these TCSs, the LPS portions of all four strains were isolated,
and the lipid-A was analyzed (Figure 6). The major signals of the WT lipid-A represent the
hexa-acylated form (m/z 1797) and
the hepta-acylated form (m/z 2036),
which is generated by the addition of a palmitate chain. Moreover,
the hydroxylation of the hexa- and hepta-acylated lipid-A could be
detected (m/z 1814, 2052). Both
modifications are catalyzed by enzymes (PagP, LpxO) whose expression
is regulated by the TCS PhoPQ.55,56 The peaks for aminoarabinose-substituted
lipid-A were not evident in the WT because this substitution (mediated
by PmrAB) occurs only under Mg2+-deficient conditions.
Thus, the lipid-A of the pmrAB-KO strain generated a similar structural
pattern as the WT in agreement with their similar susceptibility to
most of the AMPs tested here (Table 2). In comparison, the lipid-A of the phoP-KO strain
and DKO is similar and consists primarily of hexa-acylated lipid-A
(m/z 1797), which is in agreement
with their similar susceptibility to most of the AMPs (Table 2).</p><!><p>LPS analysis. MALDI-TOF
mass spectra of purified lipid-A. The major
signals represent the hepta-acylated form of lipid-A (m/z 2036) and the hexa-acylated form lacking palmitate
(m/z 1797). Moreover, the addition
of a hydroxyl group (m/z 2052) to
the hepta-acylated lipid-A could be detected in the WT and in the
pmrAB-KO.</p><!><p>Atomic force microscopy (AFM) is a powerful tool
not only for imaging the nanoscale features of bacteria but also for
measuring the mechanical characteristics of their surfaces, such as
rigidity.57,58 We explored these properties using the quantitative
nanomechanical mapping (QNM) mode of AFM. In this mode, AFM performs
multiple, fast indentations over the scanned surface, using a silicon
probe, allowing the measurement of the tip-sample interaction. The
shape of the indentation curves allows the evaluation of both the
elastic response of the bacterial surface to the AFM probe indentation
and the detachment force of the probe from the surface. It also allows
the construction of the corresponding images (maps) of the cell surface
simultaneously with regular topography images, allowing the measurement
of geometrical characteristics such as length and height (Figure 7).</p><!><p>Imaging and measurements
of bacterial cell characteristics by AFM.
(A) Three-dimensional (3D) reconstruction of S. typhimurium bacterium on a mica substrate; (B, C) DMT modulus measurement (influenced
by the thickness of the bacteria wall), measured as the mean value
over the area marked by a black rectangle; (D) measurement of the
length of the bacterium (white line), measured as the distance between
the left and right inflection points with correction for the tip-sample
convolution. (E) High-resolution image of the cell surface. Imaging
and measurements were carried out by the PeakForce quantitative nanomechanical
mapping (QNM) mode. (F) Cross section of (E) seen along the white
line.</p><!><p>In such a fashion, randomly chosen
bacterial cells were imaged,
and a flat rectangular region in their maps was chosen for the construction
of topography images (Figure 7A,C) and calculation of elastic modulus (Figure 7B). All data were normalized
to WT values. Using this analysis, we found (Figure 4) that compared to the WT, the surface of
the pmrAB-KO bacteria is much more rigid (181% of WT), whereas that
of the phoP-KO bacteria is markedly less rigid (22% of WT).</p><p>Two additional characteristics, connectivity and uniformity of
bacterial surfaces, can be explored using a "flooding analysis"51 (Figure 8). This analysis compares the height uniformity of the bacterial
surface elements. In this analysis, surface features appearing over
a certain height threshold are kept, and features beneath that threshold
are regarded as a uniform background (colored in blue). Using this
approach, a cell surface featuring no holes or shallow holes would
be regarded as uniform, whereas a surface with deep holes and dents
would be regarded as nonuniform. A complex surface would be a surface
featuring a lot of holes and elements with connectivity between them.
Visual inspection of the flooding analysis images suggests that at
a given threshold value (20 nm), WT exhibits uniform, but complex,
possibly highly interconnected, protein–protein complex structures.
This topography might be the result of the LPS and other surface elements
protruding from the surface with protein complexes and channels scattered
among them; phoP-KO bacteria, given a lower threshold value, exhibit
an almost uniform surface with a few deep holes. This can be due to
the fact that the phoP-KO membrane is thinner and more uniform, with
less protruding surface elements. This finding is further emphasized
in the TEM image of the phoP-KO bacteria (Figure 4). PmrAB-KO bacteria look more similar to
the WT and exhibit a less uniform surface and correspondingly less
complex connectivity. This might be due to tighter connections on
the surface of the bacteria. The data also reveal that the DKO bacteria
exhibit a nonuniform surface with very low connectivity, even for
a 2-fold higher threshold compared to the one used for the WT. They
also show higher ridges (lighter color). This means that the bacterial
surface has long protruding elements scattered on it, which could
be due to the uncontrolled polymerization of surface elements, as
can be seen in the TEM picture (Figure 4). These protruding elements probably prevent the tip
from penetrating into the lipidic membrane, which might give an unclear
interpretation of the bacterial surface.</p><!><p>Flooding analysis of
the bacteria using AFM. The traces shown above
are height analyses along the white lines. Sample images shown below
from the flooding analysis of WT (A), phoP-KO (B), pmrAB-KO (C), and
double-KO (D) bacterial cells. The images exhibit surface topography
above a certain height threshold. All topography features beneath
this threshold are marked blue.</p><!><p>Overall, the data reveal similar connectivity and uniformity between
the WT and PmrAB-KO in agreement with their susceptibility toward
most of the peptides tested. However, due to the uncertainty in the
data with the DKO (as explained above) and phoP-KO, it is not clear
why both of them have a similar susceptibility to most of the AMPs.</p><!><p>The PhoPQ and PmrAB TCSs have been the focus
of various studies
concerning their potential to induce bacterial resistance to AMPs.
These systems could be activated by AMPs, leading to the induction
of enzymes that modify the outer surface of Gram-negative bacteria
and make it less penetrable to AMPs. In line with that, it was shown
that the deletion of the PhoPQ system increases the susceptibility
of bacteria to AMPs.23,39 A few studies also reported the
role of PmrAB TCS in resistance toward polymyxin B and colistin, both
of which destabilize the architecture of the outer membrane through
specific interactions with LPS.23,59 However, the biophysical
aspects and the combined effects of PhoPQ and PmrAB TCSs, as well
as their contribution to bacterial resistance to cationic AMPs, are
yet not well known. We performed our study with S.
typhimurium, a frequently used model bacterium, and
investigated the roles of both PhoPQ and PmrAB systems on resistance
to a diverse array of AMPs. The results were correlated with the changes
in the biophysical properties of the bacteria. For that purpose, we
created mutants deficient in the individual systems including a DKO
mutant. The lack of these systems indeed changed the composition of
the bacterial LPS, as confirmed by mass spectrometry (Figure 6).</p><p>Using a biophysical
approach we explored the effects of the two
systems on several characteristics of the bacteria, which in turn
have an impact on the antimicrobial activity of AMPs. These characteristics
are (i) bacterial membrane polarization and permeation, (ii) surface
charge, and (iii) cell wall rigidity and density.</p><!><p>Examination of the extent of
bacterial cell polarization reveals that the phoP-KO and DKO bacteria
are similarly polarized and both are less polarized than the WT and
the pmrAB-KO (Figure 1). It has been demonstrated that the rigidities of phoP-KO and DKO
are significantly less than that of the WT, In contrast, the pmrAB-KO
bacteria are more rigid than the WT, which indicates that the phoP-KO
and DKO cell walls are more permeable than the WT and the pmrAB-KO,
which is in agreement with the higher activity of most of the peptides
on the phoP-KO bacteria compared to the pmrAB-KO.</p><p>Further support
for the differences in membrane permeability comes from the SYTOX
experiments in which we measure the extent of membrane permeability.
The data showed a trend that is similar to that of the membrane polarizing
experiment, i.e., the WT and pmrAB-KO are less permeable
than phoP-KO and the DKO (Figure 2). This is in line with changes in the composition
of the bacterial cell wall components of the different bacterial strains
due to the deletion of the TCSs, as revealed by mass spectrometry
of the LPS isolated from all of the bacteria (Figure 5). These results are also in agreement with
the MICs of most of the peptides on the different bacteria, showing
a direct correlation between permeability and susceptibility to AMPs.</p><!><p>The PmrAB
system is known to activate the pmrHFIJKLM operon, which synthesizes
and adds 4-aminoarabinose (l-Ara4N) to the LPS, making the
surface less negatively charged.60,61 Because there
is communication between the two systems through pmrD, the PhoPQ system
can also contribute to such an effect on the surface charge of the
bacteria.29,62 This effect should reduce the binding of
the positively charged AMPs to the surface, making the bacteria more
resistant. Indeed, as expected, phoP-KO and DKO bacteria bind similarly
to the pentapeptide to a higher extent than the WT bacteria. However,
the pmrAB-KO has a binding capacity similar to the WT (Figure 3), the reason for which is
not yet clear. Nevertheless, as seen with the other assays, the WT
and pmrAB-KO are similarly sensitive to most AMP. Overall, the extent
of peptide binding to the various bacteria is directly correlated
to their susceptibility to AMPs.</p><!><p>The third factor that
could contribute to the resistance of bacteria to AMPs relates to
the biophysical properties of the bacteria, which include cell wall
density and rigidity. The PhoPQ and PmrAB systems are known to induce
enzymes that add and remove fatty acids from the LPS63,64 and also modify the length of the sugar residues of the LPS.53,65 This suggests that the activation of the two systems can affect
the physical properties of the bacterial surface. To examine this
possibility, we used both TEM and AFM techniques. The TEM images of
the cross sections of the WT and the mutated bacteria shown in Figure 4 clearly reveal that
the outer membrane of pmrAB-KO bacteria is stained much stronger than
those of the WT, phoP-KO, and DKO bacteria. This suggests that the
pmrAB-KO outer surfaces are denser compared to the other three, possibly
due to the chelation of metal ions that replaced uranyl acetate during
sample preparation. This can explain why the pmrAB-KO bacteria are
not as sensitive as the phoP-KO and DKO to the AMPs investigated.</p><p>It has been revealed from the electron microscopic results that the
thickness of the periplasmic layer of the phoP-KO bacteria is significantly
smaller than that of the WT (69% of WT) and those of pmrAB-KO and
DKO to a lesser extent (86% of WT). These properties can also contribute
to the similarities in the susceptibility of phoP-KO and DKO. Regarding
pmrAB-KO, its thinner periplasmic space compared to WT can compensate
for its high rigidity, and therefore it has susceptibility to AMPs
similar to that of the WT. We also cannot rule out the possibility
that the PmrAB system affects membrane fluidity, and when it is deficient,
the membrane is denser and more rigid and thus stains stronger. The
AFM studies further demonstrated significant differences between the
rigidities of the various bacteria (Figures 7 and 8). The data
reveal that the rigidities of phoP-KO and DKO are significantly less
than that of the WT (22 and 67%, of WT, respectively). In contrast,
the pmrAB-KO bacteria are more rigid than the WT (180% of WT). This
result is consistent with our previous assays suggesting that the
lack of the PhoPQ system makes the bacterial surface less dense and
more penetrable to AMPs, while the lack of the PmrAB system makes
the surface more rigid and less penetrable.</p><p>In summary, the
PhoPQ and PmrAB TCSs have opposing effects on the
biophysical properties and biological activities of the bacteria.
The phoP-KO bacteria are more susceptible to most AMPs, have a less
rigid surface, are more penetrable, and their surface is more negatively
charged compared to WT. In contrast, the pmrAB-KO bacteria preserved
the activity of a range of AMPs, have a very dense and rigid cell
wall, and are less penetrable. This suggests that the PhoPQ system
contributes to the strengthening of the bacterial cell wall, which
helps them defend AMPs, possibly just making it more fluid and malleable.
In contrast, activation of the PmrAB system has a role in the degeneration
of the cell wall, thus making the bacteria more sensitive to AMPs.</p><p>Overall, the data suggest that the coexistence of systems with
opposing effects on the biophysical properties of the bacteria contribute
to their membrane flexibility, which is important to accommodate changing
environments but inhibits the development of meaningful resistance.</p><!><p>The results were analyzed using
a one-way analysis of variance and compared with the WT. Values of P < 0.005 were considered statistically significant.</p><!><p>Conceived
and designed the experiments: T.S., N.A.W., M.W., H.-G.S., and Y.S;
performed the experiments: T.S., M.W., A.B., and E.S; analyzed the
data: T.S., M.W., H.-G.S., and Y.S; contributed reagents/materials/analysis
tools: H.-G.S., and Y.S; wrote the paper: T.S., N.A.W., H.-G.S., and
Y.S.</p><!><p>This work was
supported in part by fundings from the German-Israel Foundation (GIF)
no. 1015-264.2/2008 (to H.-G.S. and Y.S.) and the European Community's
Seventh Framework Programme (FP7/2007-2013) under grant agreement
no. 278998 (to Y.S.). The funders had no role in the study design,
data collection and analysis, decision to publish, or preparation
of the manuscript. Y.S. is the incumbent of the Harold S. and Harriet
B. Brady Professorial Chair in Cancer Research.</p><p>The authors declare no
competing financial interest.</p>