Download The silver cation (Ag+): antistaphylococcal

J Antimicrob Chemother 2013; 68: 131 – 138
doi:10.1093/jac/dks372 Advance Access publication 25 September 2012
The silver cation (Ag1): antistaphylococcal activity, mode of action
and resistance studies
Christopher P. Randall, Linda B. Oyama, Julieanne M. Bostock, Ian Chopra and Alex J. O’Neill*
Antimicrobial Research Centre and School of Molecular and Cellular Biology, University of Leeds, Leeds LS2 9JT, UK
*Corresponding author. Tel: +44-(0)113-343-5600; Fax: +44-(0)113-343-5638; E-mail: [email protected]
Received 11 July 2012; returned 24 July 2012; revised 17 August 2012; accepted 21 August 2012
Objectives: To examine several poorly understood or contentious aspects of the antibacterial activity of silver
(Ag+), including its cidality, mode of action, the prevalence of resistance amongst clinical staphylococcal isolates and the propensity for Staphylococcus aureus to develop Ag+ resistance.
Methods: The effects of Ag+ on the viability, macromolecular synthesis and membrane integrity of S. aureus
SH1000 were assessed using established methodology. Silver nitrate MICs were determined for a collection
of staphylococcal isolates (n¼ 1006) collected from hospitals across Europe and Canada between 1997 and
2010. S. aureus biofilms were grown using the Calgary Biofilm Device. To examine the in vitro development
of staphylococcal resistance to Ag+, bacteria were subjected to continuous subculture in the presence of
sub-MIC concentrations of Ag+.
Results: Silver was bactericidal against S. aureus in buffered solution, but bacteriostatic in growth medium, and
was unable to eradicate staphylococcal biofilms in vitro. Challenge of S. aureus with Ag+ caused rapid loss of
membrane integrity and inhibition of the major macromolecular synthetic pathways. All clinical staphylococcal
isolates were susceptible to ≤16 mg/L silver nitrate and prolonged exposure (42 days) to Ag+ in vitro failed to
select resistant mutants.
Conclusions: The rapid and extensive loss of membrane integrity observed upon challenge with Ag+ suggests
that the antibacterial activity results directly from damage to the bacterial membrane. The universal susceptibility of staphylococci to Ag+, and failure to select for resistance to Ag+, suggest that silver compounds remain
a viable option for the prevention and treatment of topical staphylococcal infections.
Keywords: antimicrobial susceptibility, Staphylococcus spp., heavy metal resistance
Introduction
The silver cation (Ag+) has been used as an antimicrobial agent for
centuries and continues to be deployed for the prevention and
treatment of bacterial infection. Silver nitrate (AgNO3) is still
used in some settings as prophylaxis against ophthalmia neonatorum (neonatal conjunctivitis) and more advanced formulations
containing Ag+ are widely used to prevent and treat infection of
ulcers, chronic wounds and burns.1 – 3 In 2009, the National
Health Service spent £25 million on silver-containing dressings4
and it has been estimated that 15 metric tonnes of silver were
incorporated into medical products worldwide in 2010 alone.5 Increasingly, the antimicrobial properties of Ag+ are also being
exploited outside of the clinic, through incorporation into personal
hygiene products, textiles and water purification devices.6 – 9
Despite the long-standing and increasing use of Ag+, its antibacterial mode of action remains unclear. Work in Escherichia coli
has suggested that Ag+ exposure leads to uncoupling of the
respiratory chain from oxidative phosphorylation and loss of
the proton-motive force.10 In Staphylococcus epidermidis, exposure to Ag+ promotes release of iron from iron –sulphur clusters
and subsequent formation of the lethal hydroxyl radical, in a
process believed to result from the inhibition of electron transport chain components and production of the superoxide
anion.11 However, as Ag+ retains antibacterial activity under anaerobic conditions,12 it is unlikely that specific interaction with
the respiratory chain and formation of reactive oxygen species
is the primary mechanism by which Ag+ exerts its antibacterial
effect.
Our understanding is also incomplete regarding the potential
for development of bacterial resistance to Ag+ and the prevalence of silver resistance in clinical isolates. This may, in part,
reflect the lack of standardized methodology for assessing bacterial susceptibility to Ag+ and that surveys of Ag+ susceptibility
# The Author 2012. Published by Oxford University Press on behalf of the British Society for Antimicrobial Chemotherapy. All rights reserved.
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Randall et al.
have a small sample size and differ considerably in the methodology used,13 – 15 making it difficult to draw firm conclusions. Disparities in methodology also occur in studies to assess the
antibacterial activity of Ag+, including evaluations of cidal
potency. Consequently, AgNO3 has been reported to be rapidly
bactericidal in some studies16 and bacteriostatic in others.11
High-level resistance to Ag+ in clinically relevant Gramnegative organisms can result from the horizontal acquisition
of a plasmid harbouring the 14 kb operon encoding the Sil
protein complex.13,17 Based on homology to other heavy-metal
resistance mechanisms, the Sil system appears to confer resistance through a combination of Ag+ sequestration in the periplasm and Ag+ efflux.18,19 Resistance to Ag+ as a consequence
of chromosomal mutation has also been observed in E. coli following repeated exposure to Ag+ in the laboratory.20 Proteomic
analysis of the resulting resistant mutant revealed that expression of the native Cu+/Ag+ efflux system (CusCFBA) was upregulated when compared with the Ag+-susceptible parental
strain.21 In contrast to the situation with Gram-negative
bacteria, little is known about Ag+ resistance mechanisms in
Gram-positive bacteria.
This paper describes a series of studies to gain insights into
the issues described above. Using Staphylococcus aureus as a
model organism, we have investigated the antibacterial activity
and mode of action of Ag+ and examined the clinical prevalence
and in vitro development of silver resistance in staphylococci.
Materials and methods
Bacterial strains and culture media
The laboratory strains used in this study are described in Table 1. Clinical
staphylococcal isolates were obtained from a culture collection maintained at the University of Leeds (n¼626) and from the Chelsea and
Westminster Hospital, London, UK (n¼54), the Freeman Hospital, Newcastle, UK (n¼97), Harrogate District Hospital, Harrogate, UK (n¼82),
the London Health Sciences Centre, Victoria Hospital, Ontario, Canada
(n ¼39), the Department of Experimental Medicine and Biochemical
Sciences, University of Perugia, Italy (n ¼29) and Smith & Nephew Ltd,
York, UK (n¼79). Isolates in the University of Leeds collection were
collected worldwide (but primarily from the UK) between 1997 and
2007. Isolates from the Chelsea and Westminster Hospital, the
Freeman Hospital and Smith & Nephew were recovered between 2008
and 2010 from patients with burns or diabetic foot ulcers in clinics
where therapeutic silver was in use at the time of isolation.
Bacteria were routinely cultured using Mueller– Hinton broth and agar
(MHB and MHA, respectively; Oxoid Ltd, Cambridge, UK). For studies with
daptomycin, MHB was supplemented with Ca2+ (50 mg/L).
Antimicrobial agents, chemicals and reagents
Ciprofloxacin was from Bayer (Leverkusen, Germany), whilst daptomycin
and XF-73 were received as gifts from Cubist Pharmaceuticals (Lexington,
MA, USA) and Destiny Pharmaceuticals (Brighton, UK), respectively.
DiSC3(5) and the LIVE/DEADw BacLightTM kit were from Invitrogen Life
Technologies (Paisley, UK). The following radiolabelled chemicals were
from Perkin-Elmer (Cambridge, UK): [methyl-3H]thymidine (70 – 95 Ci/
mmol), [5,6-3H]uridine (31 –56 Ci/mmol), L-[G-3H]glutamine (20– 50 Ci/
mmol) and [1-14C]glycine. All other chemicals and antimicrobial agents
were from Sigma– Aldrich (Poole, UK). In all studies, AgNO3 was used
as the source of Ag+.
Susceptibility determinations
MICs of antibacterial agents were routinely determined by 2-fold serial
dilution in MHB according to the CLSI broth microdilution method.22
However, susceptibility of clinical staphylococcal isolates to Ag+ was
determined using the CLSI agar dilution method.22 Biofilm MICs
(bMICs) and minimum biofilm eradication concentrations (MBECs) were
determined in MHB using the Calgary Biofilm Device as described previously.23 Assessment of the cidal activity of Ag+ and comparator agents
was performed as previously described,24 using exponential-phase cultures of S. aureus SH1000 grown in MHB (108 cfu/mL) or exponentialphase cultures washed and resuspended in HEPES/glucose buffer
(5 mM HEPES, pH 7.2/5 mM glucose).
Mode of action studies
The BacLightTM assay was used to assess membrane integrity of bacteria
resuspended in water following exposure to AgNO3 and comparator
agents at 4× MIC for 10 min.25 The membrane potential of bacterial
Table 1. Bacterial strains used in this study
Species
Strain
Staphylococcus aureus
SH1000
ATCC 29213, ATCC 25923
MRSA252
Pseudomonas aeruginosa
PAO1
Escherichia coli
K12-BW25113
Bacillus subtilis
132
Description/genotype
strain 8325-4, with functional rsbU gene reinstated
S. aureus subsp. aureus Rosenbach
fully genome-sequenced EMRSA-16
Reference/source
46,47
ATCC
48
ATCC
49,50
NCTC 50110
derivative of E. coli K12 strain BD792 (lacIq rrnBT14
DlacZWJ16 hsdR514 DaraBADAH33 DrhaBADLD78)
silver-resistant E. coli K12 strain carrying plasmid pMG101
1S34
1S34 (pS72)
1S34 (pS63)
1S34 (pS77)
1S34 (pS107)
1S34 (pNS14)
parent strain of biosensors
biosensor detecting inhibition of protein biosynthesis
biosensor detecting inhibition of RNA biosynthesis
biosensor detecting inhibition of DNA biosynthesis
biosensor detecting inhibition of cell envelope biosynthesis
biosensor detecting inhibition of fatty acid biosynthesis
51
51
51
51
51
51
NCTC
JAC
Ag+ mode of action and resistance
cells resuspended in HEPES/glucose buffer was monitored using the fluorescent dye DiSC3(5).26 Leakage of K+ from cells resuspended in HEPES/
glucose buffer was determined over a 1 h time course as previously
described.26
To determine the inhibitory activity of Ag+ on the synthesis of cellular
macromolecules, the incorporation of radiolabelled precursors into DNA,
RNA, protein and peptidoglycan ([methyl-3H]thymidine, [5,6-3H]uridine,
3
14
L-[G- H]glutamine and [1- C]glycine, respectively) was monitored in
exponential-phase cultures of SH1000 (108 cfu/mL) exposed to
AgNO3 and comparator antibiotics.24,27 To assist the identification of potential cellular targets of Ag+, a collection of five Bacillus subtilis antibiotic
biosensors containing promoters specifically up-regulated by inhibitors of
DNA, RNA, fatty acid, cell envelope and protein biosynthesis were used.28
Assessment of lysis following exposure to AgNO3 and comparator
agents was carried out on exponential-phase cultures of SH1000 resuspended to an optical density at 600 nm (OD600) of 0.2 in HEPES/
glucose buffer. Following the addition of AgNO3 at 4× MIC, OD600 readings were recorded at intervals over a 120 min period. To investigate possible changes in cell morphology following exposure to AgNO3, samples
were removed at 5 and 120 min, prepared for scanning electron microscopy (SEM)29 and examined using a LEO 1530 FEGSEM scanning electron
microscope.
Selection for silver resistance
The frequency of spontaneous mutation to Ag+ resistance was determined as previously described.30 To select Ag+-resistant mutants, bacteria were continuously exposed to AgNO3 in a manner analogous to
that described by Friedman et al.31 Briefly, on day 1, broth microdilution
susceptibility testing was performed using an extended range of AgNO3
concentrations instead of the standard doubling-dilution series. Following
incubation of the cultures and determination of the MIC, the well that
contained the highest concentration of AgNO3 permitting growth was
diluted 1 :100 in MHB and used to provide the inoculum for the next
MIC assay; this process was repeated for up to 42 days.
Results
Methodology for Ag1 susceptibility testing
+
The apparent susceptibility of bacterial isolates to Ag varies
considerably depending on the culture medium used, presumably because Ag+ binds to media components such as NaCl
and proteins containing thiol groups.32,33 In addition to complicating susceptibility testing with Ag+, this effect probably also
impacts studies to investigate the cidal activity of Ag+ in
culture media. We therefore sought, in preliminary experiments,
to establish a standard set of conditions for undertaking this
work. Although originally established for the susceptibility
testing of antibiotics, we found that the CLSI agar dilution
protocol using MHA and MHB22 was appropriate for susceptibility
testing with AgNO3. Thus, this method provided high reproducibility in respect of Ag+ susceptibility across multiple determinations with a variety of staphylococcal strains (SH1000,
MRSA252, ATCC 29213 and ATCC 25923) (data not shown) and
reliably detected silver resistance (MIC .256 mg/L) in a silverresistant positive-control strain (E. coli NCTC 50110 harbouring
silver resistance plasmid pMG101). Consequently, we adopted
this method for all subsequent susceptibility testing with Ag+
and employed the same media (MHA/MHB) for our other
studies to investigate the antibacterial activity of Ag+.
Susceptibility of staphylococcal isolates to Ag1
To examine the susceptibility of clinical staphylococcal isolates to
Ag+, we conducted AgNO3 susceptibility testing as above for a
collection of 1006 staphylococcal isolates obtained from hospitals throughout Europe and Canada between 1997 and 2010.
Of these, 846 were S. aureus and 160 were coagulase-negative
staphylococci. At least 230 of these isolates came from patients
with burns and diabetic foot ulcers in clinics where therapeutic
silver was in use at the time of isolation. All isolates tested
susceptible to AgNO3 at concentrations of 8 –16 mg/L, with the
exception of a single isolate of Staphylococcus intermedius for
which AgNO3 had an MIC of 4 mg/L.
Killing kinetics of Ag1 against S. aureus SH1000
As for susceptibility testing with Ag+, there is currently no standardized method for assessing the killing kinetics of Ag+containing compounds and there are conflicting data regarding
the cidal activity of Ag+.11,16 In this study, we re-evaluated the
cidality of Ag+ using a method commonly employed for assessing the killing kinetics of antibiotics in MHB.24,27 The effect of
AgNO3 and comparator compounds (daptomycin and fusidic
acid) at 4× MIC on the viability of early exponential-phase cultures of S. aureus SH1000 grown in MHB was determined over
6 h (Figure 1a). The silver ion demonstrated only limited killing
activity compared with daptomycin, causing a ,1 log drop in
cell viability over 6 h. This result was comparable to that
obtained for fusidic acid, a bacteriostatic antistaphylococcal
agent.34 After 24 h of exposure to AgNO3, a 2.2 log drop in cell
viability was observed (data not shown). In contrast, Ag+ exhibited rapid bactericidal activity against staphylococcal cultures
resuspended in HEPES/glucose buffer, with a .7 log drop in cell
viability observed after 10 min (Figure 1b). This indicates that,
as with Ag+ susceptibility testing, the choice of assay medium
can have a significant effect on the antibacterial activity of Ag+.
Activity of Ag1 against biofilms
Bacteria growing as biofilms can cause persistent infections that
are highly refractory to antimicrobial therapy35 and may be
present in environments where silver is administered, such
as in chronic wounds.36 We therefore assessed the ability of
AgNO3 to inhibit the growth and/or eradicate biofilms of
SH1000 and MRSA252, as determined by the bMIC and MBEC, respectively. Although AgNO3 could inhibit the growth of biofilms
at a concentration equal to its planktonic MIC (bMIC 16 mg/L),
it was unable to eradicate biofilms at the highest concentration
tested (256 mg/L). The comparator agents, tetracycline and
vancomycin, were also unable to eradicate biofilm cultures at
the concentrations tested (MBEC .256 mg/L). In contrast, and
as previously reported,23 the compound XF-73 displayed potent
activity against both SH1000 and MRSA252 biofilms, with an
MBEC of 2 mg/L.
Mode of action studies with Ag1
Monitoring the ability of an antibacterial compound to inhibit the
biosynthesis of cellular macromolecules can provide insights into
133
Randall et al.
8
8
7
7
6
6
Log10 cfu/mL
(b) 9
Log10 cfu/mL
(a) 9
5
4
3
5
4
3
2
2
AgNO3
Untreated control
Daptomycin
1
AgNO3
1
Fusidic acid
0
0
0
100
200
Time (min)
300
400
0
10
20
30
Time (min)
40
50
60
Figure 1. Effects of AgNO3 and comparator agents on the viability of exponential cultures of S. aureus SH1000 over a 6 h period in MHB (a) and over a
1 h period in cultures resuspended in HEPES/glucose buffer (b). Values shown are the means of three replicates from three independent experiments.
Error bars represent standard deviations from the mean.
134
100
DNA
Protein
Peptidoglycan
RNA
90
80
Incorporation (%)
70
60
50
40
30
20
10
at
e
ac
yc
lin
Si
e
lv
er
ni
tr
at
Va
e
nc
om
yc
Si
in
lv
er
ni
tr
at
e
Ri
fa
m
pi
ci
Si
n
lv
er
ni
tr
at
e
tr
tr
ni
Te
er
Si
lv
pr
o
flo
xa
ci
n
0
Ci
its mode of action.24,26,27 We therefore assessed the ability of
AgNO3 to inhibit the incorporation of radiolabelled precursors
into DNA, RNA, protein and peptidoglycan, and compared this
with the activity of antibacterial agents known to specifically
inhibit these biosynthetic pathways (ciprofloxacin, rifampicin,
tetracycline and vancomycin, respectively). At 4× MIC, AgNO3
caused substantial and comparable inhibition of all four biosynthetic pathways after 10 min (Figure 2).
To further explore the antibacterial target of Ag+, we employed
five B. subtilis antibiotic biosensors, each containing a luciferase reporter gene fused to promoters induced by inhibitors of fatty acid,
protein, DNA, RNA and cell envelope biosynthesis. These constructs
have previously been shown to be of benefit in mode-of-action
studies of antibacterial compounds.28 Exposure of the strains to
AgNO3 failed to elicit a significant increase in luminescence in
any of the biosensors (data not shown).
The non-specific inhibition of biosynthetic processes by AgNO3
may result from damage to the cell membrane and consequent
loss of intracellular components.24 Thus, following exposure of
S. aureus SH1000 to AgNO3 and comparator antibiotics at 4×
MIC, we assessed bacterial membrane integrity using the
BacLightTM assay and membrane potential using the fluorescent
dye DiSC3(5). Following 10 min of exposure to Ag+, the membrane integrity decreased by 97%. This decrease was comparable to that seen for bacteria exposed to nisin (99%), a
compound known to damage bacterial membranes through
pore formation.37 The membrane potential was also affected
Figure 2. Effects of AgNO3 and comparator agents on DNA, RNA, protein
and peptidoglycan biosynthesis in S. aureus SH1000, as measured
by incorporation of radiolabelled precursors. Values represent the
percentage incorporation relative to drug-free controls. Values shown
are the means of three replicates from three independent experiments.
Error bars represent standard deviations from the mean.
JAC
Ag+ mode of action and resistance
(a)
(b)
100
100
90
80
80
Potassium remaining (%)
Membrane potential (%)
90
70
60
50
40
70
60
50
40
Nisin
30
30
AgNO3
20
20
Untreated control
10
10
0
0
0
10
20
30
Time (min)
40
50
60
0
10
20
30
Time (min)
40
50
60
Figure 3. Effect of AgNO3 and nisin on the membrane potential of S. aureus SH1000 (a) and on the ability of S. aureus SH1000 to retain K+ (b). Values
shown are the means of three replicates from at least three independent experiments. Error bars represent standard deviations from the mean.
in cells exposed to Ag+, with a .50% decrease observed over 1 h
(Figure 3a). The kinetics of membrane damage following Ag+
exposure were further explored by measuring the leakage of
intracellular K+ from S. aureus SH1000 over 1 h (Figure 3b). In
,10 min, .90% of intracellular K+ was lost from cells exposed
to Ag+; again, this effect was comparable to that observed for
bacteria exposed to nisin.
Cells treated with nisin eventually undergo lysis as a consequence of damage to the membrane.37 As Ag+ exposure produced similar results to nisin in the BacLightTM and K+ leakage
assays, we examined whether Ag+ also causes cell lysis.
S. aureus SH1000 cells resuspended in HEPES/glucose were
exposed to AgNO3 at 4× MIC and monitored for lysis over a
period of 2 h by optical density measurements. The results
were compared with those of nisin-exposed cultures (Figure 4).
Over 2 h, the optical density of the cultures exposed to nisin
decreased by 37%, indicating that lysis was occurring. There
was no change in the optical density of cultures exposed to
Ag+
suggesting that under these conditions silver did not
,
cause lysis.
To confirm the absence of lysis and to detect any changes in
the morphology of cells exposed to Ag+, cells were visualized
using SEM following exposure to Ag+ or nisin for 5 and
120 min (Figure 4). Cells visualized after 120 min of exposure
to nisin showed evidence of lysis, but no lysis was seen for
cells treated with AgNO3 (Figure 4).
Selection of bacterial Ag1 resistance
Spontaneous mutants of S. aureus SH1000 with reduced susceptibility to Ag+ could not be obtained (limit of detection
,1×10210) following plating of saturated cultures onto MHA
containing AgNO3 at 4× MIC. In contrast, spontaneous mutation
to resistance to the comparator agents mupirocin and fusidic
acid occurred at frequencies of 5×1028 and 3×1026, respectively, consistent with previously reported values.38,39 Failure to
obtain spontaneous mutants exhibiting reduced susceptibility
to Ag+ led us to adopt an alternative method to attempt
to select for resistance. S. aureus SH1000 and the clinical
S. aureus strain MRSA252 were continuously exposed to AgNO3
in the form of repeated microdilution broth MICs.31 Attempts
were also made to select Ag+ resistance in Pseudomonas aeruginosa PAO1 and E. coli K12-BW25113, in order to determine
whether the rate at which Ag+ resistance develops differs in
Gram-negative organisms. Following 6 days of exposure, the
AgNO3 MIC for E. coli increased from 4 to .256 mg/L, while
AgNO3 MICs for S. aureus and P. aeruginosa cultures remained
at 16 and 4 mg/L, respectively, for the duration of the experiment (42 days). In contrast, S. aureus cultures exposed to daptomycin developed clinically significant levels of daptomycin
resistance (.2 mg/L) following 3 days of exposure (data not
shown).
Discussion
In this study, the CLSI agar dilution susceptibility testing method
was used to determine Ag+ MICs for 1006 staphylococcal isolates, using AgNO3 as the source of Ag+. Since there are no published breakpoints for Ag+, the Ag+ MIC that corresponds to
clinically significant Ag+ resistance is unknown.40 However, as
the MICs of Ag+ for the isolates in this study were comparable
to those for S. aureus SH1000 and other laboratory strains, no
135
Randall et al.
0.350
0.300
0.250
OD600
0.200
0.150
0.100
Nisin
AgNO3
0.050
0.000
0
20
40
60
Time (min)
80
100
120
Figure 4. Effects of Ag+ and nisin on the integrity of S. aureus SH1000 resuspended in HEPES/glucose buffer. Values shown are the means of three
replicates from three independent experiments. Broken lines point to scanning electron micrographs of cells taken from cultures at indicated
timepoints. Error bars represent standard deviations from the mean.
evidence of reduced susceptibility to silver in clinical isolates of
staphylococci was found. The universal susceptibility of staphylococci to Ag+ suggests that silver therapy could be considered as
an alternative for the treatment of topical staphylococcal skin
infections that are resistant to commonly used topical antibiotics. Furthermore, as this method is based on CLSI guidelines
commonly used for antibiotic susceptibility testing, it could
easily be established in clinical laboratories to allow for continued monitoring of Ag+ susceptibility.
Studies in the recent scientific literature have employed
various methodologies to assess the killing kinetics of
Ag-containing compounds. Differences include the choice of
culture medium, the starting inoculum size and the concentration of the Ag-containing compound used.11,16 This has led to
conflicting results regarding the cidality of Ag+. Using a
method frequently employed for assessing the bactericidal activity of antibiotics in susceptibility-testing media (MHB), Ag+ was
found to exhibit bacteriostatic activity against S. aureus
SH1000, causing only a 2.2 log10 drop in cell viability over 24 h.
However, in buffer, Ag+ was rapidly bactericidal, causing a
.7 log10 drop in cell viability in 10 min. The considerably
increased cidal potency of Ag+ in buffer is presumably primarily
due to the lack of NaCl and thiol-containing proteinaceous components that are present in culture media, which have been
reported to bind and inactivate Ag+.32,33 These results emphasize
the importance of adopting standardized culture media (such as
MHB) when assessing the antibacterial activity of Ag+-containing
compounds.
Exposure of S. aureus to Ag+ caused rapid loss of membrane
integrity. Since assays to measure bacterial membrane integrity
require that bacterial cells are resuspended in buffer (or water),
136
in conditions under which Ag+ is potently bactericidal, care
must be taken when relating these findings to the antibacterial
mode of action of Ag+ in growth media. However, when bacterial
cells were exposed to Ag+ for 10 min in MHB, simultaneous inhibition of four major macromolecular biosynthesis pathways
was observed. Such non-specific inhibition has previously been
observed for compounds that cause membrane damage and is
likely a result of leakage of cytoplasmic constituents from the
cell and subsequent de-energization of biosynthetic pathways.24,26 As this process occurs rapidly upon addition of Ag+
to cultures and correlates with the results of membrane
damage assays in buffer, it seems likely that the cell membrane
is indeed the primary target of Ag+.
The mechanism by which Ag+ damages the bacterial membrane is unclear. Unlike cells exposed to nisin, no lysis or
changes in morphology were observed for cells exposed to Ag+.
The direct interaction of Ag+ with thiol-containing macromolecules in the membrane, or hydroxyl radical production as a consequence of Ag+ inhibiting electron transport, may contribute to the
observed membrane damage.11 However, since Ag+ maintains
activity (albeit at a reduced level) in anaerobic conditions,12 hydroxyl radical production cannot be the sole cause of membrane
damage. Future studies will focus on understanding how Ag+ acts
to compromise the integrity of the bacterial membrane.
It has been suggested that antibacterial agents that target the
bacterial membrane may be beneficial for the treatment of persistent infections, such as those caused by non-growing bacteria
or those resident in biofilms,41,42 since such membrane-active
agents typically retain antibacterial activity against metabolically
inert bacteria.23,41 We therefore sought to determine the antibacterial activity of Ag+ against biofilm cultures of SH1000 and
JAC
Ag+ mode of action and resistance
MRSA252. At a concentration of 16 mg/L, AgNO3 could prevent
shedding of planktonic cells from established biofilms in vitro.
This may indicate that, in vivo, Ag+ could prevent further biofilm
growth and dissemination to other sites. However, as this bMIC
is equivalent to the MIC of Ag+, it seems likely that Ag+ inhibits
the growth of planktonic cells following their release from the
biofilm and has no direct effect on the viability of the biofilm
itself. Indeed, we determined that Ag+ was unable to eradicate
established biofilms (MBEC .256 mg/L). This may be a consequence of Ag+ binding to negatively charged extracellular
polymeric substances within the biofilm,43 thereby preventing
Ag+ from reaching cells in a sufficient concentration to exert an
antibacterial effect.
The continuous exposure of cells to AgNO3 for 42 days failed
to select for Ag+ resistance in S. aureus or P. aeruginosa. The inability to select endogenous bacterial Ag+ resistance highlights
another potential advantage of using Ag+ in place of antibiotics
for the treatment of topical staphylococcal infections. Although
we could not generate Ag+-resistant S. aureus or P. aeruginosa
in this study, we were able to select E. coli resistant to
.256 mg/L AgNO3 following a short period of continuous subculture. The generation of Ag+-resistant E. coli in the laboratory
has previously been reported and appears to result from the
increased expression of the CusCFBA efflux transporter.20,21
However, as E. coli is rarely isolated from wounds where therapeutic Ag+ is used,44,45 it remains unclear if Ag+-resistant
E. coli would arise or persist in the clinical setting. It is worth
noting that neither S. aureus nor P. aeruginosa harbour homologues of the CusCFBA system, which may provide an explanation
for the inability to select for Ag+ resistance in these species.
In summary, our work suggests that standardization of
culture media and experimental methodology are critical when
evaluating the antibacterial activity of Ag+. Although no overt
lysis can be observed following exposure to Ag+, the rapid and
extensive loss of membrane integrity strongly suggests that
the antibacterial activity of Ag+ results from direct damage to
the cell membrane. The universal susceptibility of staphylococcal
isolates to Ag+, coupled with the inability to select for Ag+ resistance following repeated exposure, suggest potential advantages
of silver-based therapy over antibiotics for the treatment and
prevention of superficial staphylococcal infections.
Acknowledgements
A part of this work was presented at the Twenty-first European Congress
of Clinical Microbiology and Infectious Diseases and Twenty-seventh
International Congress on Chemotherapy, Milan, Italy, 2011 (C. Randall,
L. Oyama, J. Bostock, I. Chopra and A. J. O’Neill; Poster P1454).
We thank N. Mughal (Chelsea and Westminster Hospital, UK), D. Tate
(Freeman Hospital, UK), Z. Hussain (London Health Sciences Centre,
Canada), A. Mencacci (University of Perugia, Italy) and Smith &
Nephew for provision of staphylococcal isolates. All trademarks are
acknowledged.
Funding
This work was supported by an MRC DTG-CASE studentship with Smith
& Nephew.
Transparency declarations
C. P. R. has received travel grants from Smith & Nephew to attend international scientific meetings. I. C. has acted as a consultant to Smith &
Nephew and has received research grants from the company. All other
authors: none to declare.
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