University of Groningen
Responses of Staphylococcus aureus to mechanical and chemical stresses
Carniello, Vera
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57
Chapter 3
Surface Enhanced Fluorescence and Nanoscopic Cell Wall Deformation in Adhering Staphylococcus aureus Upon Exposure to Cell Wall Active and Non-Active Antibiotics
Vera Carniello, Brandon W. Peterson, Jelmer Sjollema, Henk J. Busscher, Henny C. van der Mei
Nanoscale (2018), 10:11123-11133 Reprinted with permission from Royal Society of Chemistry
Chapter
ABSTRACT
In infection, bacteria often adhere to surfaces and become deformed by the forces with which they adhere. Nanoscopic cell wall deformation defines bacterial respons-es to environmental conditions and is likely influenced by antibiotics. Here, staphylo-coccal cell wall deformation upon exposure to cell wall active and non-active antibi-otics or their combinations, are compared for two green-fluorescent (GFP) isogenic
Staphylococcus aureus strains adhering to a gold surface, of which one lacks
pep-tidoglycan cross-linking. Exposure to cell wall active antibiotics caused greater cell wall deformation than a buffer control in the GFP parent and in the Δpbp4GFP isogenic
mutant, as measured by surface-enhanced-fluorescence. Cell wall non-active antibi-otics only yielded greater deformation than a buffer control in the parent strain, while combinations of cell wall active and non-active antibiotics did not cause greater cell wall deformation. 3D-analysis of the impact of adhesion forces and Young’s mod-uli of the cell wall, both measured using atomic force microscopy, yielded the con-clusion that increased deformation was mainly due to cell wall weakening and not due to effects of antibiotics on adhesion forces. Interactions between bacteria and antibiotics are mostly studied using planktonic bacteria, while in infection bacteria are in an adhering state, that deforms their cell wall and therewith influences their adaptive responses. We anticipate that the demonstration of cell wall weakening in adhering bacteria under the influence of antibiotics and the role of peptidoglycan herein, will aid the development of new antibiotics. Surface-enhanced-fluorescence may accordingly develop into a new, highly-sensitive method for diagnosing anti- biotic-resistant bacteria.
59 INTRODUCTION
Bacterial cell walls are composed of a rigid, cross-linked peptidoglycan layer en-compassing the lipid membrane holding the cytoplasm, and a softer outer layer consisting of different types of proteinaceous surface appendages and extracellu-lar polymeric substances (EPS). The cell wall provides protection against osmotic pressure and maintains shape and structure of bacterial cells [1]. Cell wall integrity therewith plays a crucial role in bacterial survival, as under antibiotic attack. Several antibiotics, such as vancomycin, are cell wall active and disrupt cell wall integrity or inhibit its synthesis [2,3]. Other cell wall active antibiotics, like oxacillin, bind to cell wall components to trigger release of autolytic enzymes causing bacterial cell lysis [4,5]. Antibiotics non-active on cell walls, penetrate through the cell wall into the cytoplasm with the aim of preventing RNA or protein synthesis [6,7]. In clinical microbiology, cell wall active and non-active antibiotics can be used in concert to control infection.
In infection, bacteria often adhere either to each other, to tissue cells, calcified tis-sues like bone and teeth or to biomaterials implants and devices [8]. Accordingly, during antibiotic treatment of infection, bacteria are not only exposed to the chemi-cal stress of antibiotics, but also to a mechanichemi-cal stress arising from nanoscopic cell wall deformation under the influence of the forces by which they adhere. Simultane-ous exposure of Staphylococcus aureus to the antibiotic nisin and strong adhesion to polyethylene yielded enhanced activation of an intra-membrane located sensor pro-tein that caused greater efflux of nisin than could be achieved by planktonic staph-ylococci not adhering to a surface [9]. Bacterial cell walls possess many different types of sensor molecules to probe their environment. Cell wall deformation, with its impact on the lipid membrane, is an important factor defining the bacterial response to its environment, including adhesion to surfaces. Therefore, better understanding of cell wall deformation mechanisms and factors that influence it, like possibly cell wall activity of antibiotics, may lead to new strategies to enhance antimicrobial ef-fectiveness. Moreover, nanoscopic bacterial membrane deformation is considered as a new, sensitive option for diagnosing antibiotic-resistant bacteria [10].
Opposite to the large deformation of tissue cells upon adhesion to a surface, that can be readily quantified using simple light or electron microscopic techniques[11], bacterial cell wall deformation is nanoscopically small due to the rigidity provided to the bacterial cell wall by its peptidoglycan layer. Reliable measurement of nano-scopic bacterial cell wall deformation in fluorescent bacterial strains has become possible by exploiting surface enhanced fluorescence (SEF) [12]. SEF is the increase
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in fluorescence that occurs when a fluorophore approaches a reflective metal sur-face. Due to a combination of surface plasmon resonance and a mirror effect, the fluorescence of the fluorophore-metal complex can be enhanced up to hundreds of times the fluorescence of the fluorophore alone [13–16]. The effect of SEF decreas-es exponentially with the distance of the fluorophore from the surface and is only detectable for fluorophores within distances up to about 30 nm from a metal surface [17]. Upon adhesion of a fluorescent bacterium to a metal surface, nanoscopic cell wall deformation under the influence of adhesion forces will gradually increase and bring more fluorophores within the cytoplasm closer to the metal surface within the range of SEF. Accordingly, the degree of SEF has been demonstrated to be propor-tional with cell wall deformation [12].
The aim of this study is to compare cell wall deformation upon exposure to cell wall active and non-active antibiotics, using SEF of two green-fluorescent isogenic
S. aureus strains adhering to a gold surface, of which one lacks cross-linking of its
peptidoglycan. Atomic force microscopy (AFM) was applied to quantitate the ad-hesion forces of the strains to the gold substratum on which the experiments were conducted and their Young’s moduli.
MATERIALS AND METHODS
Bacterial strains, growth conditions and antibiotics
Green-fluorescent (GFP) S. aureus ATCC 12600GFP and its isogenic Δpbp4GFP mutant,
with GFP located in the cytoplasm, were grown on Tryptone Soya Broth (TSB; OX-OID, Basingstoke, England) agar plates supplemented with 10 μg mL-1 tetracycline
(Sigma-Aldrich, St. Louis, MO, USA) at 37 °C for 24 h. Next an overnight culture (TSB, supplemented with tetracycline) prepared from a single colony, was diluted 1:20 in 100 mL of fresh TSB without tetracycline supplementation and incubated at 37 °C for 16 h.
Cultures were harvested by centrifugation (5000 × g) and washed twice in phos-phate buffer (5 mM potassium phosphos-phate [2.5 mM K2HPO4, 2.5 mM KH2PO4], 0.075 M NaCl, pH 7.0), supplemented with 3 % v/v TSB to maintain bacterial viabil-ity in suspension. The bacterial suspension was then sonicated (3 × 10 s, 30 W) in ice-water bath (Vibra Cell Model 375, Sonics and Materials Inc., Danbury, CT, USA). The bacterial concentration was determined with a Bürker-Türk counting chamber and the suspension was diluted in phosphate buffer supplemented with TSB to a concentration of 3 × 108 mL-1.
61 For antibiotics, two cell wall active antibiotics [3,4], vancomycin hydrochloride (Xel-lia Pharmaceuticals ApS, Copenhagen, Denmark) and oxacillin sodium salt (Sigma- Aldrich, St. Louis, MO, USA), and two cell wall non-active [6,7], rifampicin (Sigma- Aldrich St. Louis, MO, USA) and gentamicin sulfate salt (Sigma-Aldrich St. Louis, MO, USA), were dissolved in phosphate buffer supplemented with TSB.
Staphylococcal cell surface properties
Contact angles on bacterial lawns and surface free energies
The hydrophobicity of the staphylococcal cell surfaces was determined through contact angle measurements with different liquids on staphylococcal lawns using the sessile drop technique and a home-made contour monitor. Briefly, bacteria were deposited on a 0.45 μm pore-size HA membrane filter (Millipore Corporation, Bedford, MA, USA) using negative pressure. Filters were then dried until reaching constant, so-called “plateau” water contact angles, representing removal of free water from the lawn, while leaving bacterial cell surfaces in a hydrated state [18]. Subsequently, contact angles were measured with ultrapure water, formamide and α-bromonaphthalene on three different lawns from different bacterial cultures. These liquids have different polarities (Table S1), allowing to calculate the electron-donating (γ-) and accepting
(γ+) parameters of the acid-base (γAB) surface free energy component which,
togeth-er with the Lifshitz-Van dtogeth-er Waals (γLW) component, constitute the total surface free
energy (γtot) [19].
Zeta potentials
In order to compare the charge properties of the bacterial cell surfaces, staphylococ-ci were resuspended in 10 mM potassium phosphate buffer at different pH values (pH 2, 3, 4, 5, 7) and particulate microelectrophoresis (Zetasizer Nano ZS, Malvern Instruments, Worcestershire, United Kingdom) was carried out to measure the elec-trophoretic mobilities, from which zeta potentials were derived employing the Helm-holtz-Smoluchowski equation [20]. The experiment was performed in triplicate with different bacterial cultures.
Microbial adhesion to hydrocarbons (kinetic MATH assay)
The combined effects of surface hydrophobicity and charge with respect to the ad-hesion of the staphylococci to a hydrophobic ligand were determined, as previously described [21]. Briefly, bacteria were resuspended to an optical density at 600 nm be-tween 0.4 and 0.6 (initial absorbance at time zero [A0]) in 3 mL of 10 mM potassium phosphate buffer at different pH values (pH 2, 3, 4, 5, 7) containing 1:20 hexadecane as photospectrometrically measured (Spectronic 20 Genesys, Spectronic Instru-ments, Rochester, NY, USA). After vortexing the suspension for 10 s and settling for
Chapter
10 min, the optical density was measured again (absorbance at time t [At]), and this procedure, according to the kinetic MATH protocol, was repeated for five more times to allow calculation of an initial rate of bacterial removal from the aqueous phase (Figure 1A) according to
(1) where t is the vortexing time. The experiment was performed in triplicate with differ-ent bacterial cultures.
Antibiotic effects on staphylococcal strains
Minimal inhibitory and minimal bactericidal concentrations
Bacterial cultures (106 mL-1 in TSB) were dispensed into each well of a 96-well
micro-titer plate with known antibiotic concentrations in TSB, with a step factor dilution of 2 starting from 256 μg mL-1, and incubated at 37 °C for 24 h. Following incubation, the
minimal inhibitory concentration (MIC) was taken as the lowest antibiotic concentration not creating visible turbidity. Then, 10 μL of bacterial suspensions of each well showing no turbidity were plated on TSB agar plates and incubated at 37 °C for 24 h. The mini-mal bactericidal concentration (MBC) was taken as the lowest concentration at which no colonies were visible on the plate. The experiment was performed in triplicate with different bacterial cultures.
Time-kill kinetics
Staphylococcal cultures (3 × 108 mL-1 in phosphate buffer supplemented with TSB)
were diluted 1:10 in antibiotic solutions in phosphate buffer supplemented with TSB. After 0, 0.5, 1, 1.5, 2 h, suspensions were serially diluted in PBS (10 mM potassium phosphate, 0.15 M NaCl, pH 7.0) and 100 μL aliquots were plated on TSB agar plates and incubated for 24 h at 37 °C. The number of colonies formed on the plate was then manually counted.
Surface enhanced fluorescence
For SEF, staphylococci suspended in phosphate buffer supplemented with TSB were injected in a parallel plate flow chamber (PPFC), having borosilicate glass (Men-zel-Gläser, Menzel GmbH&Co KG, Braunschweig, Germany) top and bottom plates in absence of flow to allow complete sedimentation of all staphylococci. Complete sedimentation was verified by enumeration of the number of bacteria on the bot-tom plate using phase-contrast microscopy (107 bacteria cm-2 on average) and
com-parison with the number of staphylococci injected. The bottom plate was coated with 10 nm gold (DLRI, St. Charles, MO, USA), possessing a water contact angle of Roboto light:
(1)
TFE(t) =R(0) - RR(t) - R0 0
(2)
Rate of initial removal= limt → 0 dt log $d AAt
0 x 100%
(3)
63 80 ± 1 degrees. A Teflon spacer separated the top and bottom plates, yielding a channel height of 0.058 cm. Gold-coated glass slides were used as received, after rinsing with demineralized water.
Next, fluorescence imaging was performed with a bio-optical imaging system (IVIS Lumina II, Caliper LifeScience, Hopkinton, MA, USA), with an excitation wavelength of 465 nm and emission in a range from 515 to 575 nm. Fluorescence images had a field of view 5 × 5 cm, exposure time of 90 s and temperature during imaging was 20 °C. Images were displayed on a pseudo-color scale overlapped to a grey-scale im-age, to demonstrate homogeneous distribution of bacterial fluorescence in the flow chamber (Figure S1). The Living Image software package 3.1 (Caliper LifeScience) provided a user-defined region of interest (ROI, about 5 × 1.2 cm) for each image and was used to calculate the average fluorescence radiance (photons s-1 cm-2
steradi-an-1) over the user-defined ROI.
For measurements, background fluorescence in the user-defined ROI was first mea-sured in the PPFC filled with phosphate buffer supplemented with TSB and subtract-ed from all fluorescence intensities measursubtract-ed. Next, bacteria were injectsubtract-ed into the PPFC and allowed to sediment under the influence of gravity while acquiring images every 15 min for 3 h, sufficient for complete sedimentation of all bacteria in suspen-sion [12]. Subsequently, the PPFC was filled with antibiotic solution and fluorescent image acquisition continued every 15 min for an additional 2 h. Antibiotic concentra-tions applied were based on results of MIC and MBC experiments (see Table 1), and amounted 0.004 μg mL-1 for rifampicin, 1 μg mL-1 for gentamicin, 0.125 μg mL-1 for
oxacillin and 1 μg mL-1 for vancomycin, while phosphate buffer supplemented TSB
was used as buffer control. For these antibiotic concentrations, antibiotic exposure did not affect bacterial fluorescence, as established in a separate experiment (data not shown) and also supported in similar experiments [22].
Assuming even distribution of GFP in the cytoplasm, the increase in fluorescent radiance due to adhering staphylococci relative to planktonic bacteria was expressed as total fluorescence enhancement (TFE) [12]
(2) where R(t) is the fluorescence radiance at time t, R(0) is the fluorescence radiance measured for a planktonic suspension and R0 is the fluorescence radiance of the background. Note that a major advantage of SEF above other methods to measure cell wall deformation, is that individual bacteria do not need to be imaged and defor-Roboto light:
(1)
TFE(t) =R(t) - R0 R(0) - R0
(2)
Rate of initial removal= limt → 0 dt log $d AAt
0 x 100%
(3)
F - Fadh= 43 E* &R d3
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mation is calculated from an enhanced fluorescence over a large field of view. Pho-tobleaching was negligible under these experimental conditions, as established in a separate experiment (data not shown) and as supported by literature [12]. All SEF experiments were performed in triplicate with different bacterial cultures.
Atomic force microscopy (AFM)
Single bacterial-contact AFM probes were prepared by immobilizing bacteria on NP-O10 tipless cantilevers (Bruker, Camarillo, CA, USA), as described previously [23]. Briefly, at the onset of each experiment, cantilevers were calibrated by the thermal tuning method yielding spring constants in the range 0.03 - 0.12 N m-1 and mounted on a micromanipulator (Narishige International, Tokyo, Japan) under microscopic observation (Leica DMIL, Wetzlar, Germany). The apex of the cantilever was sub-sequently dipped into a droplet of 0.01 % poly-L-lysine (molecular weight 70,000 to 150,000, Sigma-Aldrich) for 1 min, dried in air for 2 min and immersed into a droplet of bacterial suspension for 2 min. Immobilization of S. aureus on AFM cantilevers functionalized with poly-L-lysine had no negative effect on bacterial viability [24,25]. Imaging a calibration grid (HS-20MG BudgetSensors, Innovative Solutions Bulgar-ia Ltd., SofBulgar-ia, BulgarBulgar-ia) with the bacterBulgar-ial probe confirmed single-bacterBulgar-ial contact with the surface [9], and probes yielding double contour lines were discarded (which seldom or never happened). Fluorescence microscopy images of bacterial probes confirmed that bacteria were always positioned at the centerline-front of the apex of the cantilever. Positioning at the lateral edge of the apex would provide force curve shapes different from regular bacterial probes. In that case, the probe was discarded and replaced by a new one.
Prior to AFM measurements, antibiotics were added to a bacterial suspension (3 × 108 mL-1 in phosphate buffer supplemented with TSB) and incubated for 2 h
at room temperature. Centrifugation (5000 × g) was applied to separate bacteria from the antibiotic solution, and staphylococci were subsequently resuspended in 10 mM potassium phosphate buffer (pH 7.0, 106 bacteria mL-1) and sonicated. AFM
force measurements were done at room temperature in 10 mM potassium phos-phate buffer (pH 7.0) on a BioScope Catalyst atomic force microscope (Bruker), us-ing gold-coated glass slides as substratum surfaces. Force-distance curves were recorded under a loading force of 3 nN, at an approach and retraction velocity of 2 μm s-1, and bond-maturation times of 0, 2, 5 and 10 s were applied. To verify that
measurements did not disrupt bacterial integrity, five force-distance curves at a loading force of 3 nN and 0 s contact time were acquired at the beginning and end of each experiment. When adhesion forces measured differed more than 1 nN from the beginning to the end of an experiment, the probe was discarded and replaced
65 by a new one. For each strain and antibiotic, AFM measurements were performed with three probes prepared from three different bacterial cultures. With each probe, three different spots on the gold-coated glass slide were measured, recording five force-distance curves in each spot for each contact time.
Reduced Young’s moduli E* of the staphylococci were obtained by the Derjagu-in-Muller-Toporov (DMT) fit of the contact region of the retraction force-distance curves
(3) where F is the force exerted by the cantilever on the bacterium, Fadh is the adhesion force, R the radius of the bacterium, assumed to be 500 nm for both strains [23], and d the bacterial deformation.
Statistical analysis
GraphPad Prism, version 5 (San Diego, CA, USA) was employed for statistical anal-ysis. Data were tested for normal distribution with Shapiro-Wilk normality test. If data were normally distributed, a two-tailed Student’s t-test was employed. When data were not normally distributed as for the Young’s moduli, non-parametric Mann– Whitney U-test replaced the Student’s t-test, respectively. p < 0.05 was used as sig-nificance for all tests.
Roboto light: (1)
TFE(t) =R(0) - RR(t) - R0 0
(2)
Rate of initial removal= limt → 0 dt log $d AAt
0 x 100%
(3)
F - Fadh= 43 E* &R d3
Chapter
Figure 1. Physico-chemical surface properties of the strains employed. (A) Schematic presentation of the kinetic MATH assay.
(B) Initial bacterial removal rates of staphylococci from an aqueous phase (10 mM potassium phosphate buffer) by hexadecane as a function of pH. Error bars represent standard errors over measurements on three different bacterial cultures.
(C) Zeta potentials of staphylococci in 10 mM potassium phosphate buffer as a function of pH. Error bars represent standard errors over measurements on three different bacterial cultures. (D) Contact angles on staphylococcal lawns with ultrapure water (θw), formamide (θf), α-bro-monaphthalene (θb) and bacterial surface free energy parameters and components. γ- and
γ+ are the electron-donating and electron-accepting parameters, while γAB and γLW are the
acid-base and Lifshitz-Van der Waals components of the total surface free energy, γtot. ± signs
represent standard errors over measurements on three bacterial lawns, prepared from three different bacterial cultures. Averages are not signifi cantly different between the two strains (Student’s t-test, p > 0.05).
67 RESULTS
Physico-chemical surface properties and antibiotic susceptibility of the strains First, physico-chemical surface properties relating to adhesion of S. aureus ATCC 12600GFP and its isogenic mutant S. aureus ATCC 12600 Δpbp4GFP lacking
cross-link-ing of its peptidoglycan, were evaluated. Both strains showed low initial removal rates from an aqueous phase by hexadecane (Figures 1A and 1B) that were slightly higher for the parent strain. Low removal by hexadecane is either indicative of a hydrophilic cell surface, the presence of electrostatic double-layer interactions be-tween bacteria and hexadecane droplets or a combination of both [26]. Therefore, staphylococcal zeta potentials were measured as a function of pH (Figure 1C). Zeta potentials were negative over the entire pH trajectory from pH 2 to 7, while being slightly less negative for the parent strain. Water contact angles on staphylococcal lawns deposited on membrane filters were less than 60 degrees for both strains, while being somewhat, but not statistically significant (p > 0.05), lower for the parent strain. Also α-bromonaphthalene contact angles differed slightly but not significantly (p > 0.05) between both strains, while formamide contact angles were similar (Figure 1D). In a surface thermodynamic analysis, these differences in contact angles trans-lated in a slightly, but not significantly (p > 0.05) higher electron-donating surface free energy parameter for the parent strain as compared to the Δpbp4GFP mutant,
minor differences in electron-accepting parameters and Lifshitz-Van der Waals com-ponents, finally resulting in a slightly, but not significantly higher total surface free energy (p > 0.05) for the parent strain (see also Figure 1D). Collectively, these data demonstrate that the outermost cell surfaces of both strains are highly similar with extremely minor differences that bear no biological significance. Accordingly, the isogenic mutant can be considered to differ from its parent strain in peptidoglycan cross-linking but not in outermost cell surface properties.
Minimal inhibitory (MIC) and bactericidal (MBC) concentrations for two cell wall non-active and active antibiotics are summarized in Table 1. In order to carry out SEF experiments on metabolically active cells, it was decided on the basis of Table 1 to carry out experiments on cell wall weakening under the influence of antibiotics at a concentration equal to the lowest MIC across the two strains for each antibiotic. Accordingly, it could be demonstrated that both strains remained viable, i.e. losing far less than one log-unit of colony forming units (CFUs), upon exposure to either of the cell wall non-active or active antibiotics or combinations of cell wall active and non-active antibiotics (Figure 2).
Chapter
Table 1. Minimal inhibitory (MIC) and minimal bactericidal (MBC) concentrations and mode of action for the different antibiotics and bacterial strains. Data are averages of triplicate experi-ments with different bacterial cultures.
S. aureus ATCC 12600GFP S. aureus ATCC 12600 Δpbp4GFP
Cell wall non-active antibiotics
Rifampicin (RIF) MIC (μg mL-1) 0.004 0.004 MBC (μg mL-1) 0.5 0.25 Gentamicin (GEN) MIC (μg mL-1) 4 1 MBC (μg mL-1) 4 1
Cell wall active antibiotics
Oxacillin (OXA) MIC (μg mL-1) 0.25 0.125 MBC (μg mL-1) 16 0.25 Vancomycin (VAN) MIC (μg mL-1) 1 2 MBC (μg mL-1) 8 16
Figure 2. Bacterial survival in presence of (sub)-MIC of antibiotics. Log (CFU) per mL in staphylococcal suspensions as a function of time in the presence and absence of different cell wall non-active (RIF, rifampicin and GEN, gentamicin) and cell wall active (OXA, oxacillin and VAN, vancomycin) or combinations thereof. Data represent single-fold measurements.
69 Fluorescence enhancement
Total fluorescence enhancement (TFE; see Figure 3A) increased rapidly up to 2 h af-ter staphylococcal adhesion to gold surfaces, mostly due to bacaf-terial sedimentation, and leveled off for the parent strain at a stationary level of 1.36 ± 0.02, while for the isogenic mutant deficient in peptidoglycan cross-linking, TFE reached a significantly higher (p < 0.0005) level of 1.66 ± 0.04 after 3 h, confirming the relative weakness and greater deformability of its cell wall (compare Figures 3B and 3C). Continued ex-posure to buffer after injection of buffer as a control after 3 h, did not yield a further increase in TFE for the parent strain (Figure 3D), but the isogenic Δpbp4GFP mutant
demonstrated ongoing cell wall deformation. Exposure to antibiotics of the adhering staphylococci after 3 h of adhesion to the gold surface yielded a further increase in TFE, that was quantified as an area under the curve (AUC; see inset to Figure 3B) with respect to the stationary level of TFE in buffer observed after 3 h for the strain under consideration (Figure 3D). Note that the AUC after continued exposure to the buffer control, was significantly (p < 0.05) lower for the parent strain than for the Δpbp4GFP isogenic mutant due to ongoing deformation of the mutant strain in buffer
(Figure 3E). Subsequently, the AUC data for the different cell wall non-active and active antibiotics and their combinations, were averaged. Cell wall non-active anti-biotics only yielded greater cell wall deformation than the buffer control in the par-ent strain (see also Figure 3E), while cell wall active antibiotics caused significantly (p < 0.05) greater cell wall deformation than the buffer control both in the parent strain as well as in the Δpbp4GFP isogenic mutant. Combinations of cell wall active
and non-active antibiotics with respect to the buffer control did not cause great-er cell wall deformation. Alike the buffgreat-er control, cell wall active antibiotics or their combinations with cell wall non-active antibiotics caused greater deformation in the Δpbp4GFP isogenic mutant than in the parent strain (p > 0.05).
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Figure 3. Cell wall deformation of adhering staphylococci upon antibiotic exposure.
(A) Schematic presentation of SEF for measuring nanoscopic bacterial cell wall deformation. Upon adhesion of a fluorescent bacterium to a metal surface from its planktonic state, cell wall deformation will gradually increase under the influence of adhesion forces exerted by the substratum and bring more fluorophores within the cytoplasm closer to the metal sur-face within the range of sursur-face enhancement. Since the attractive molecular forces arising from the substratum act pairwise on all molecules of the organisms including their cell wall, these forces yield nanoscopic deformation bringing more fluorescent molecules in the SEF range at equal intracellular volume. This is exactly the same deformation process that causes spreading and severe deformation, in fact visible flattening, in mammalian cells adhering on a surface. Deformation in mammalian cells however, is not counteracted by a rigid layer, such as the peptidoglycan layer in bacteria, and is therefore microscopic instead of nanoscopic. (B) Total fluorescence enhancement (TFE) as a function of time for S. aureus ATCC 12600GFP.
Antibiotics were added 3 h after initiating staphylococcal sedimentation and adhesion on a gold surface. Inset represents the area under the curve (AUC) after introduction of antibiotics (in green) with respect to the stationary level in TFE observed after 3 h in buffer. Note that the AUC could also have been obtained from the fluorescence levels of each strain with respect to the buffer value at each point in time, but this option was not preferred as it would not allow us to directly compare S. aureus ATCC 12600 Δpbp4GFP with its parent strain. Although it would
have yielded different AUC values, the conclusions of the work would not be affected. (C) Same as (B), for S. aureus ATCC 12600 Δpbp4GFP.
(D) Increases in total fluorescence enhancement expressed as AUC (see Figures 3B and C) upon exposure of staphylococci adhering to the gold surface to different cell wall non-active (RIF, rifampicin and GEN, gentamicin) and cell wall active (OXA, oxacillin and VAN, vancomycin)
71 Adhesion forces and bacterial cell wall elasticity
Next, the adhesion forces between the adhering staphylococci and the gold surface, responsible for the cell wall deformations observed, were measured as a function of the contact-time between the bacteria and the substratum surface using single bac-terial probe AFM (see Figures 4A and 4B). Force-distance curves (see Figure 4C for an example) demonstrated a clear minimum force when the distance between the interacting surfaces increased, which was taken as the “adhesion force”. In Figure 4D it can be seen that upon initial contact, i.e. after 0 s bond maturation, both the parent strain and the Δpbp4GFP isogenic mutant had a similarly low adhesion force
of 2.4 ± 0.4 and 1.7 ± 0.6 nN respectively, that increased differentially with contact time. These increases in adhesion forces with contact time could be fitted to an exponentially increasing function, allowing calculation of stationary adhesion forces and characteristic time constants for bond maturation. Stationary adhesion forces were significantly (p < 0.0005) smaller for the Δpbp4GFP isogenic mutant than for the
parent strain (Figure 4E and 4F), regardless of whether exposed to cell wall non-ac-tive or acnon-ac-tive antibiotics or their combinations. Both in the parent strain and in the Δpbp4GFP mutant, exposure to cell wall active or non-active antibiotics or their
combi-nations yielded small difference in stationary adhesion forces, that ranged from 10.6 to 13.6 nN and from 2.9 to 4.1 nN in the parent strain and the mutant, respectively (Figure 4F). Regardless of whether exposed to a buffer control, cell wall non-active or active antibiotics, the parent strain, with its more rigid cell wall, achieved stationary adhesion forces significantly more slowly (characteristic bond maturation time 4.0 ± 0.2 s on average) than the mutant strain (2.5 ± 0.3 s). Exposure of the Δpbp4GFP
mutant to combinations of antibiotics increased the bond maturation time to the same level as of the parent strain (4.4 ± 0.5 s versus 3.9 ± 0.3 s, respectively). Since cell wall deformation depends on an interplay between adhesion forces and cell wall elasticity, reduced Young’s moduli were obtained by fitting the linear region of the retraction force-distance curves, i.e. where the bacterium is still in contact with the surface, according to the Derjaguin-Muller-Toporov (DMT) model. Young’s moduli decreased significantly (p < 0.05) after continued exposure to antibiotics as antibiotics or combinations thereof. Error bars represent standard errors over three experi-ments with different bacterial cultures.
(E) Same as in Figure 3D, but now with the increases in total fluorescence enhancement av-eraged over each category or combination of antibiotics. Error bars represent standard errors combining all six replicates with cell wall active or non-active antibiotics, and 12 replicates with antibiotic combinations. Significant differences between groups are indicated (Student’s t-test, p < 0.05).
Chapter
compared with continued exposure to a buffer control (Figure 4G) for both strains. Averaging data for the different types of antibiotics or their combination revealed a signifi cantly (p < 0.05) lower Young’s modulus for the Δpbp4GFP mutant compared
to the parent strain (Figure 4H). Young’s moduli for the parent strain after antibiotic exposure did not differ signifi cantly (p > 0.05) for antibiotics with different modes of action, while for the Δpbp4GFP mutant, Young’s moduli were reduced to different
73
Figure 4. Atomic force microscopy on staphylococci upon antibiotic exposure. (A) Schematic presentation of single bacterial-contact AFM probe.
(B) Fluorescence image of a single S. aureus ATCC 12600GFP on the apex of the cantilever. Scale
bar represents 15 µm.
(C) Examples of force-distance curves for S. aureus ATCC 12600 Δpbp4GFP on a gold surface
after bond maturation for different contact-times.
(D) Examples of adhesion forces as a function of bond maturation time for staphylococci on gold exposed to buffer. Dashed lines indicate an exponential fi t, allowing to derive a stationary adhesion force and characteristic bond maturation time constant. Error bars represent stan-dard errors over 45 force-distance curves obtained on 9 different spots on the gold surface and three probes prepared from three different bacterial cultures.
(E) Stationary adhesion forces of staphylococci after 2 h of antibiotic exposure. Error bars rep-resent standard errors over 45 force-distance curves obtained on 9 different spots on the gold surface and three probes prepared from three different bacterial cultures.
(F) Same as (E), but now with adhesion forces averaged over each category or combination of antibiotics. Error bars represent standard errors combining all 90 force-distance curves with cell wall active or non-active antibiotics, and 180 force-distance curves with antibiotic combinations. Signifi cant differences between groups are indicated (Student’s t-test, p < 0.05). (G) Reduced Young’s moduli of adhering staphylococci after 2 h of antibiotic exposure. Whis-kers represent the minimum and maximum values, boxes represent the 25th–75th percentile, and the solid horizontal line indicates the median value over 45 force-distance curves obtained on 9 different spots on the gold surface and three probes prepared from three different bac-terial cultures.
(H) Same as (G), but now with Young’s moduli averaged over each category or combination of antibiotics. In total, averaging involved 90 force-distance curves with cell wall active or non-ac-tive antibiotics, and 180 force-distance curves with antibiotic combinations. Signifi cant differ-ences between groups are indicated (Mann–Whitney U-test, p < 0.05).
Chapter
DISCUSSION
In this paper, differential cell wall deformation has been observed in staphylococ-ci adhering to a gold surface upon exposure to cell wall active and non-active an-tibiotics using SEF (Figure 3A). Both a buffer control, cell wall active anan-tibiotics as well as their combinations with cell wall non-active antibiotics caused greater deformation in the Δpbp4GFP isogenic mutant than in the parent strain (p > 0.05).
Both types of antibiotics yielded greater deformation in an adhering S. aureus strain with a cross-linked, rigid peptidoglycan layer with respect to a buffer control. In a Δpbp4GFP isogenic mutant with a weakened cell wall, only cell wall active antibiotics
yielded greater deformation than the buffer control, while cell wall non-active anti-biotics caused similar deformation. Combinations of cell wall active and non-active antibiotics did not cause greater cell wall deformation.
Cell wall deformation results as an interplay between adhesion forces sensed by a bacterium and the cell wall rigidity. However, adhesion forces are not entirely inde-pendent on cell wall deformation, as a deformed bacterium has a larger contact area with the substratum which invokes a stronger adhesion force. Since neither bacterial zeta potentials, nor surface energetics differ between the parent and the Δpbp4GFP
isogenic mutant strain, and effects on staphylococcal removal by hexadecane are absent (Figure 1), it can be concluded that the outermost cell surface in both staph-ylococcal strains is very similar and the Δpbp4 mutation involves cross-linking of the peptidoglycan located in the deeper cell wall without affecting the outermost cell surface. This is reflected in the adhesion force measurements, that show an identi-cally low adhesion force directly upon contact of around 2 nN for the parent strain and the mutant, respectively. Upon bond maturation however, deeper cell wall lay-ers other than the outermost cell surface, may become involved in adhesion, which eventually yielded a stronger stationary adhesion force for the rigid parent strain as compared with the more deformable, Δpbp4GFP isogenic mutant. Intriguingly, the
rigid parent strain will possess a smaller contact area than the mutant strain, yet a larger adhesion force was observed. Hence the Δpbp4GFP mutation, though not
af-fecting the outermost cell surface, must not only affect peptidoglycan cross-linking but also deeper cell wall components that become involved in adhesion upon bond maturation [27]. Said differently, a cell wall is not a 2D surface, but a very complex, 3D structure, with many layers including outermost surface molecules and appendages, fuzzy coats and a deeper-located peptidoglycan layer and lipid membrane. Bacterial cell surface characterization as done here, proves that the outermost cell surface is identical in both bacterial parent and mutant strains, showing that the difference in cross-linking of the peptidoglycan of the Δpbp4 mutant must be confined to and
75 located in the deeper cell wall, and is not affecting the outermost cell surface. During bond maturation, the outermost cell surface re-arranges and deeper parts of the cell wall may become exposed towards the surface, affecting adhesion forces between a substratum and bacterial cell surface. For instance, the mutation lowers the pro-duction of poly-N-acetylglucosamine (PNAG) [28], which prevents the development of strong binding with the substratum surface. These suggestions are in line with the observation that in the Δpbp4GFP mutant initial adhesion forces increase much faster
to a stationary state than in the parent strain, in which a higher stationary adhesion force is achieved.
From a clinical perspective, the behavior of the parent strain is most useful to dis-cuss. Most antibiotics, regardless of their cell wall activity, or combinations yield larger deformation with little or no significant differences in adhesion forces, hence differences in deformation must be ascribed to differences in cell wall rigidity. Cell wall active antibiotics can reduce bacterial cell wall elasticity by a variety of mecha-nisms. Exposure of Escherichia coli to a high concentration of cell wall active ampi-cillin dissolved in phosphate buffered saline (PBS) provoked not only lysis and leak-age of cytoplasmic content [29], but also inhibited peptidoglycan synthesis and fast changes in transcription profile [30], accompanied by small molecular movements in the cell wall [31]. Cell wall active antibiotics like vancomycin have also been sug-gested to insert themselves into the cell wall to increase the local mechanical strain, i.e. the local deformation, and weaken the cell wall [32], as in the growth of Borrelia
burgdorferi in the presence of sub-inhibitory concentrations of vancomycin [33]. In
addition, a smaller periplasmic space between the cell wall and the outer membrane was observed, while growth at vancomycin concentrations higher than MIC generat-ed bacteria with “blebbing” of the cell wall in a dose-dependent manner, presumably due to weakening of the cell wall [33].
Cell wall non-active antibiotics are taken up intra-cellularly through antibiotic solu-bilization in the lipid membrane or through pores formed by porin proteins [34] or mechanosensitive channels [35]. The current effects of cell wall non-active antibi-otics on cell wall rigidity suggest that these antibiantibi-otics also cause cell wall weaken-ing in naturally occurrweaken-ing bacteria. Such effects cannot be a priori ruled out as the transport of cell wall non-active antibiotics to the cytoplasm necessitates crossing the rigid peptidoglycan layer, explaining possibly why the MIC of cell wall non-active antibiotics of the Δpbp4GFP was 4 times lower than of the parent strain. Other studies,
evaluating the effects of antibiotics included in growth medium, have reported that cell wall non-active tobramycin, another aminoglycoside like gentamycin, inhibits the synthesis of the enzymes responsible for peptidoglycan synthesis [36,37]. Also
sim-Chapter
ilar experiments in Pseudomonas aeruginosa have demonstrated that growth in the presence of sub-inhibitory concentrations of tobramycin induces the production of abnormal cell wall proteins which incorporate themselves in the bacterial cell wall, making it lose integrity and reduce its elasticity, as has been demonstrated using AFM [36,37]. Our study, though carried out under very different experimental con-ditions, suggest that the cell wall non-active antibiotics rifampicin and gentamicin and their combinations with cell wall active antibiotics may have similar effects in
S. aureus.
The role of adhesion forces in establishing cell wall deformation is unexpectedly small compared to the role of the cell wall’s Young’s modulus. In the 3D presentation of cell wall deformation as a function of adhesion forces and Young’s moduli (Figure 5), it can be seen that over the force range from 5 – 15 nN only a minor change in deformation occurs. Note that in many adhesion phenomena such as adhesion to inert surfaces or bacterial (co-)aggregation, differences in bacterial adhesion forces of several nNs have already been described as influential [38,39], which emphasizes the relatively small role of adhesion force in deformation. Oppositely, the entire range of staphylococcal deformations observed can be largely explained by changes in Young’s moduli, representing cell wall weakening.
CONCLUSIONS
Exposure of S. aureus adhering to a gold surface to cell wall active antibiotics caused greater cell wall deformation than a buffer control in strains with and without cross-linked cell wall peptidoglycan. Cell wall non-active antibiotics only yielded greater de-formation than a buffer control in the parent strain. The role of differences in bacteri-al adhesion forces in cell wbacteri-all deformation after exposure to different antibiotics was small, yielding the conclusion that increased deformation upon antibiotic exposure was solely due to cell wall weakening and not due to effects of antibiotics on adhe-sion forces. Interactions between bacteria and antibiotics are mostly studied using planktonic bacteria, while in infection bacteria are in an adhering state, that deforms their cell wall and therewith influences their adaptive responses, including emergent properties like EPS production that may protect the organisms against antibiotics [40]. We anticipate that the demonstration of cell wall weakening in adhering bacte-ria under the influence of antibiotics and the role of peptidoglycan herein, will aid the development of new antibiotics. Surface enhanced fluorescence may accordingly develop into a much needed [10], new, highly-sensitive method for diagnosing anti-biotic-resistant bacteria.
77 CONFLICTS OF INTEREST
HJB is also a director of a consulting company, SASA BV. We declare no potential conflicts of interest with respect to authorship and/or publication of this article. The opinions and assertions contained herein are those of the authors and are not con-strued as necessarily representing the views of the funding organization or the au-thors’ employers.
ACKNOWLEDGEMENTS
This study was entirely funded by the University Medical Center Groningen, Gronin-gen, The Netherlands. We are grateful to Mariana G. Pinho, Laboratory of Bacterial Cell Biology, and Sergio R. Filipe, Laboratory of Bacterial Cell Surfaces and Pathogen-esis, Instituto de Tecnologia Quimica e Biológica, Universidade Nova de Lisboa, for providing the pMAD-pbp4 plasmid.
Figure 5. The interplay of adhesion force and Young’s modulus determining bacterial cell wall deformation upon adhesion to a surface. Data points are averages for each strain and an-tibiotic or combination thereof, as included in this study. The plane is obtained by orthogonal linear regression of all data points in the three-dimensional space. Color bar represents the magnitude of AUC. Squares represent data points above the regression plane, triangles are data points below the plane. Data in blue represent S. aureus ATCC 12600GFP, while red data
points represent S. aureus ATCC 12600 Δpbp4GFP. Color scale bar is artificial and not related to
strain indication.
Chapter
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81 SUPPLEMENTARY MATERIAL
Liquid Water Formamide α-bromonaphthalene
γ- 25.5 39.6 0
γ+ 25.5 2.3 0
γAB 51.0 19.0 0
γLW 21.8 39.0 44.4
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Table S1. Surface free energy parameters and components of the three liquids used for con-tact angle measurements. γ- and γ+ are the electron-donating and electron-accepting
parame-ters, while γAB and γLW are the acid-base and Lifshitz-Van der Waals components, respectively.
All data in (mJ m-2) and taken from Van Oss et al. [1]
Figure S1. Representative fluorescence image of sedimented, fluorescent S. aureus ATCC 12600GFP in a parallel plate flow chamber, as used for SEF experiments, showing
homoge-neous fluorescence distribution in the center of the chamber, which was taken as the ROI in SEF. Scale bar is 1 cm. Pseudo-color bar represents the photon count.
