Atomic Force Microscopy Reveals the Mechanobiology of Lytic

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Atomic force microscopy reveals the mechanobiology of lytic peptide action on bacteria Anna Mularski, Jonathan J. Wilksch, Huabin Wang, Mohammed Akhter Hossain, John D. Wade, Frances Separovic, Richard A. Strugnell, and Michelle L Gee Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.5b01011 • Publication Date (Web): 15 May 2015 Downloaded from http://pubs.acs.org on May 17, 2015

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Atomic force microscopy reveals the mechanobiology of lytic peptide action on bacteria

Anna Mularski1, Jonathan J. Wilksch2, Huabin Wang1, Mohammed Akhter Hossain1,3, John D. Wade1,3, Frances Separovic1, Richard A. Strugnell2*, Michelle L. Gee1

1

School of Chemistry, The University of Melbourne, VIC 3010 Australia

2

Department of Microbiology and Immunology, The Peter Doherty Institute for Infection

and Immunity, The University of Melbourne, VIC 3010, Australia 3

Florey Institute for Neuroscience and Mental Health, The University of Melbourne, VIC

3010, Australia. *Corresponding author

Keywords: atomic force microscopy, Klebsiella pneumoniae, antimicrobial peptide, capsular polysaccharide, membrane interaction

ACS Paragon Plus Environment

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Abstract

Increasing rates of antimicrobial-resistant medically-important bacteria require the development of new, effective therapeutics, of which antimicrobial peptides (AMPs) are amongst the promising candidates. Many AMPs are membrane-active but their mode of action in killing bacteria or in inhibiting their growth, remains elusive. This study used atomic force microscopy (AFM) to probe the mechanobiology of a model AMP (a derivative of melittin) on living Klebsiella pneumoniae bacterial cells. We performed in situ biophysical measurements to understand how the melittin peptide modulates various biophysical behaviours of individual bacteria, including the turgor pressure, cell wall elasticity, and bacterial capsule thickness and organisation. Exposure of K. pneumoniae to the peptide had a significant effect on the turgor pressure and Young’s modulus of the cell wall. The turgor pressure increased upon peptide addition followed by a later decrease, suggesting that cell lysis occurred and pressure was lost through destruction of the cell envelope. The Young’s modulus also increased, indicating that interaction with the peptide increased the rigidity of the cell wall. Surprisingly, the bacterial capsule did not prevent cell lysis by the peptide, and the capsule appeared unaffected by exposure to the peptide, as capsule thickness and inferred organisation was within the control limits, determined by mechanical measurements. These data show that AFM-measured turgor pressure may provide valuable insights into the physical events that precede bacterial lysis by AMPs.


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Introduction

Bacterial resistance to antibiotics is increasing at an alarming rate, threatening to dramatically impact our ability to treat common infections.

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This rise in resistance, coupled with

relatively low numbers of new antibiotics approved by regulatory bodies such as the US Food and Drug Administration, underscores the importance of researching and developing new therapeutic agents to treat bacterial infections.3 Antimicrobial peptides (AMPs) are considered to be a promising alternative to conventional antibiotics.

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These membrane

active peptides are amphipathic, short chain and cationic, and found throughout the plant and animal kingdoms, forming an important part of many hosts’ innate defence response against bacterial infection. The minimum inhibitory concentrations (MICs) of AMPs are in the low micromolar range, similar to the active concentrations of currently-used antibiotics against sensitive organisms. 7

The mechanism of how AMPs interact with the bacterial membrane and induce cell lysis is currently unclear. Some studies point to a pore-forming mechanism

8-10

whereas others

suggest detergent-like action.11-12 The majority of these studies have been conducted using synthetic model membrane systems often isolated from the dynamic processes that continually remodel the bacterial surface and cell wall13-14. Recently however, studies have revealed that AMP interactions with both model membrane systems and live bacteria are not always analogous15-17. The work of Gee et al. using fluorescence lifetime imaging suggests that pore formation is a less significant part of lipid-peptide interaction in live bacteria than in model membrane systems.


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Atomic Force Microscopy (AFM) is a powerful technique to study the surfaces of biological samples in situ18-19. To date, investigating AMP bacterial interactions using AFM has typically been done in air due to relative ease of imaging 20-21. The AFM images obtained in these studies reveal changes in cell height and surface roughness over time. Unfortunately, when imaging in air, the dehydration of biological samples provides opportunities for artefacts to develop, and be observed. These previous studies of AMPS have revealed cell damage as a result of AMP treatment, but the damage caused by the AMPs is difficult to separate from the damage caused by the air-drying, even when control samples are used. Although experimentally more complex, AFM imaging of biological samples in fluids revealed less dramatic changes in cell height and surface roughness following treatment with an AMP. 22-23

More recently, AFM has proved to be useful for measuring physical properties and intermolecular forces on live biological cells under non-destructive, physiologically relevant conditions that require little sample preparation24-27. Some of these studies have specifically looked at how AMPs impact adhesion and cell stiffness in live bacteria28-30. Gaboriaud et al. 31

, Wang et al.

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, have demonstrated that properties such as long-range electrostatic

interaction, cell wall elasticity and cytoplasmic turgor pressure of bacteria can be assessed using AFM.

In this present study, we used AFM to interrogate the physiological responses of the bacterium, Klebsiella pneumoniae, when exposed to a model AMP. K. pneumoniae are recognized as a serious cause of nosocomial infections and a source of shared antimicrobial resistance such as the extended-spectrum beta lactamases (ESBLs) and, more recently, carbapenem-resistance33. K. pneumoniae typically assemble a polysaccharide layer (known as


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the capsule) on the outside of the cell envelope. It takes the form of a hydrated polyelectrolyte network which can grow up to several hundred nanometers thick,

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and can

provide protection from environmental stresses, including desiccation, detergents, antibiotics and host immune defenses.

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The exopolysaccharide capsule plays a role in bacterial

adherence to surfaces (such as medical devices) through the formation of biofilms.36 Llobet et al.

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used purified capsular polysaccharides to increase the resistance of a capsule-deficient

K. pneumoniae mutant to polymyxin B and α-defensin-1. The purified capsular polysaccharides increased the MICs of the AMPs against a range of bacteria, leading to the suggestion that the bacterial capsule could act as an effective bacterial countermeasure to resist AMPs.

The role of the capsule in mediating AMP-cell interactions has been interrogated here using the AFM methodologies developed by Gaboriaud et al.

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and Wang et al.

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where the

Pincus equation is used to probe the organisation and thickness of the K. pneuomiae capsule. The Young’s modulus, derived from the Hertz equation, gives a measure of envelope stiffness or elasticity and Hooke’s law allows for the determination of the cytoplasmic turgor pressure or the pressure the bacterial cytoplasm, plasma membrane and peptidoglycan exert against the bacterial outer membrane. In this paper, we present AFM nanomechanical force data in situ of live K. pneumoniae when exposed to a model antimicrobial peptide that is a derivative of melittin, the main toxin of honey bee (Apis mellifera) venom.

Melittin is one of the most well characterised membrane-lytic peptides10, 13, 38-39 and is a 26 residue peptide (GIGAVLKVLTTGLPALISWIKRKRQQ-NH2) which adopts an alpha helical structure in membrane environments. We show that the peptide had a significant effect on the turgor pressure and Young’s modulus of the cell wall. The observed increase


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and subsequent decrease in turgor pressure suggests that cell lysis occurred due to peptide accumulation at the membrane and that cytoplasm escaped relatively slowly through a compromised cell envelope. The Young’s modulus also increased, indicating that interaction with the peptide increased the stiffness of the cell wall. The bacterial capsule did not prevent cell lysis by the lytic peptide, nor did the capsule appear to be affected by exposure to the peptide, as the thickness and organisation varied no more than as for a control population of bacteria, indicating that ionic interaction of bacteria-bound capsular polysaccharide is not a key factor for AMP interaction with live K. pneumoniae cells.


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Experimental

1. Peptide synthesis The melittin mutant (GIGAVLKVLTTGLKALISWIKRKRQQ-NH2) used in this study, melittin (P14K), was formed by substituting Pro-14 with Lys. This substitution allowed amine coupling of the fluorescent label AlexaFluor 430 for allowing future spectroscopic work on this system. The solid phase synthesis of the peptide was described previously by Rapson et al.

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Characterisation by Rapson et al. revealed the labelled mutant peptide has

significantly greater lytic capability than native melittin in artificial membrane systems.

2. Bacterial strains, culture conditions, and harvesting Klebsiella pneumoniae AJ218 (capsule serotype K54) is a human urinary tract infection isolate, identified at the microbiological laboratory of the Alfred Hospital, Melbourne, Australia.

41-42

All strains were maintained on Luria-Bertani (LB) agar at 37°C. LB broths

inoculated with these cultures were grown for 16 hours at 37°C while shaking (180 rpm). Stationary phase cells were then harvested by centrifugation (10 min at 3500 × g) and washed twice with Milli-Q™ water (18.2 MΩ cm-1). The final concentration of bacterial cells in Milli-Q™ water was approximately 2 × 108 CFU mL-1.

A wzc mutant of K. pneumoniae AJ218, defective in the transporter that enables capsule polysaccharide export, was isolated following random mini-Tn5Km2 transposon insertion mutagenesis43 of K. pneumoniae AJ218. To confirm transposon insertion within the wzc gene, Y-linker ligation PCR and subsequent DNA sequencing analysis of the transposon flanking region was performed to ensure correct location of the mutation. 44


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The minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC) of melittin was determined for both wild-type and capsule-deficient K. pneumoniae cells using the microdilution broth method outlined by the National Committee for Clinical Laboratory standards.45

3. Bacterial sample conditions for AFM measurements Polyethyleneimine (PEI) coated glass slides were used to immobilize wild-type bacteria. Capsule-deficient bacteria were immobilised on gelatin-coated glass slides as capsuledeficient K. pneumoniae AJ218 did not adhere firmly enough to PEI-coated slides for force measurements. The PEI and gelatine coating methods are described by Wang et al. 32

Substrate rigidity is a requirement when measuring cell indentation to ensure that only cell compression contributes to the measurement. Wang et al.

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have shown that, even though

gelatin may be expected to be a softer substrate than PEI, there is no measureable effect of gelatin deformability on the force profiles of the bacteria, which suggests that this method results in a gelatin layer thin enough to present as a rigid surface in AFM force measurements.

Bacteria were immobilised by adhering onto PEI or gelatin coated surfaces. All mechanical measurements were performed within 2-3 hours of removal of the bacteria from growth media. Bacteria-coated slides were immersed in 10 mM HEPES buffer (pH 7.4) or 1.2 µM peptide solution (made in 10 mM HEPES buffer, pH 7.4) and kept stationary within the calibrated AFM for at least 40 minutes before measurements commenced. This methodology precludes shaking of bacterial samples prior to measurement, but allows for immediate imaging and force profile collection after microscope equilibration is completed. Under


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comparable conditions, cells remained viable for the duration of the experiment as determined by live/dead cell fluorescence assays using a Molecular Probes Bacterial Viability Kit.

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When the fluorescence assay was conducted on peptide treated cells, 30 minutes after

treatment with 1.2 µM peptide solution, ~82% of cells had compromised membranes (Supporting Information, Figure S2).

An imaging volume of 5 mL was used. Measurements were conducted first in HEPES buffer and then exchanged for 1.2 µM peptide in HEPES buffer. Preliminary experiments for this work were performed at considerably higher concentrations of peptide, but imaging proved difficult and measurement of forces curves was not possible. The peptide concentration was successively lowered until imaging and force curve collection was unambiguous and reproducible. To establish the lipid to peptide ratio at which experiments were conducted, bacteria attached to the slides were stained with crystal violet following AFM measurements, and viewed at 60x magnification. Counting cells in several areas of known size allowed for an approximation of the number of cells per slide. Work reported by Ingraham et al.

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gives

an approximation of the number of lipids per microbial cell, which allowed us to calculate of the number of lipid molecules per slide. The lipid to peptide ratio for all experiments reported here was estimated to be 1:30. This ratio was selected since it is considerably higher that that generally used in experiments on model membrane systems to ensure measurable peptide-cell interaction within the timeframe of the experiments.

4. Atomic force microscopy and force measurements AFM measurements were performed using an MFP-3D instrument (Asylum Research, Santa Barbara, CA). Silicon nitride cantilevers were purchased from Bruker (MLCT, Camarillo, CA) with a nominal spring constant of 0.01 Nm-1 and probe radius of 20 nm (according to


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manufacture specifications). Cantilever spring constants were determined using the thermal tune method 47 included in the MFP-3D software. Calibrated spring constants were within the range of 0.016-0.020 Nm-1. All cantilevers used were from the same batch. All tips were cleaned in a BioForce UV/ozone cleaner (BioForce Nanosciences Inc, Ames, IA) before use. Photodetector sensitivity was measured on a clean silica slide prior to force measurements. 48 The slope of the constant compliance region of the force curves obtained was used to convert the deflection, d, in mV to nm. The cantilever defection was then converted into a force, F, according to Hooke’s law, F = k × d, where k is the force constant of the cantilever. 49

Cells were imaged in contact mode at a scan rate of 1 Hz. Trace and retrace were monitored to locate the true apex of cells and ten force curves were measured at different locations along it. The cell apex was probed during force measurements rather than the cell periphery, which has a high degree of curvature that makes quantifying mechanical properties difficult. 50

Imaging was repeated after each collection of ten force curves to ensure no change in cell

morphology had occurred. Force curves were acquired at a loading rate of 600 nm s-1.

Force data at time zero (i.e., before the addition of peptide) were acquired by measuring 10 force curves per cell across a population of cells. The values obtained were averaged to provide a negative control and are in good agreement with the data reported by Wang et al. 32 in similar conditions. Average values for turgor pressure and Young’s modulus of cells in buffer are shown in Figure 3. Zero time values for polymer brush length and capsule thickness are not shown in Figure 5 as these parameters vary from cell to cell, typically 98 (±13) nm and 315 (±46), respectively, for untreated cells and what is more relevant is the change in value with time.


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The force profiles acquired following the addition of peptide were measured on single wildtype cells over a two-hour time frame. Capsule-deficient cells did not adhere to the gelatincoated surface with sufficient strength to withstand multiple force profile collections. As such, force curves of capsule-deficient cells were collected across a population of cells. At each time point, 10 force curves were measured and averaged. Experiments were performed five times to ensure reproducibility in the observed trends over time. The error bars around each data point in the figures below represent ±1 standard deviation, which was observed to be similar at each point in time. The data obtained showed good reproducibility from location to location along each cell apex and from cell to cell at comparable times of exposure to peptide (Supporting Information, Figure S3).

5. Cell viability assays Cell viability assays were performed using a LIVE/DEAD BacLight Bacterial Viability Kit (Molecular Probes, Eugene OR). Wild-type and capsule-deficient K. pneumoniae cells were adhered to modified glass slides that were prepared in the same way as for AFM experiments (described above). The purpose was twofold: to confirm that the bacteria were viable at the commencement of an AFM experiment, and to determine the rate at which cells were compromised after exposure to the peptide. Cells were visualised and images captured using a Zeiss LSM 700 confocal microscope and Zeiss Zen imaging software. Green and red channel images were thresholded using ImageJ. Cells were counted using the ‘Analyse particles’ function. Microscopy was performed at the Biological Optical Microscopy Platform, University of Melbourne.


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Results and Discussion

Bacterial cells adhered to either PEI or gelatine-coated slides were imaged in contact mode in HEPES buffer (pH 7.4) before and after force measurements. Figure 1a shows a typical 12 x 12 µm AFM image of well-dispersed K. pneumoniae cells. The bacterial cells are rod-shaped, as expected, and the surfaces of the cells appear topographically homogenous on this scale and smaller scale images (Figure 1b).

1μm

0

-1

0.7μm

0

-0.7

Figure 1. 3D projections of AFM images: (a) 12 × 12 µm image of topographically homogenous, rod shaped K. pneumoniae cells; and (b) 5 × 5 µm image of a single K. pneumoniae cell. All cells were imaged in situ (10 mM HEPES buffer, pH 7.4).


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Nanomechanical force profiles for the indentation of bacterial cells were obtained at ten points along the apex of each bacterium to provide statistically robust data. Lateral drift was minimised by allowing the system to equilibrate before commencing measurements and the absence of drift was confirmed by comparing images of cells before and after force measurements. Force curves collected in this way were highly reproducible (Supporting Information, Figure S3). The maximum loading force was controlled in all experiments to avoid puncturing the bacterial cell wall with the AFM tip.

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Note that the contact point i.e.

zero indentation, is defined as the point at which contact was first made between the tip and the sample, as measured by the onset of cantilever deflection. Long-range non-contact double layer interactions are small in this study since the electrolyte concentration is 10 mM, at which the expected Debye length is ~3 nm.

Figure 2(a) shows a typical force profile for the indentation of a K. pneumoniae cell in HEPES buffer. Wang et al. 32 have demonstrated that the indentation of K. pneumoniae cells is comprised of four distinct stages when working at low electrolyte concentrations. The authors have developed a mechanical model for cell indentation (summarised in Supporting Information, Figure S1) where different regions of each force curve are fitted using the model that best describes the cell surface-AFM tip interaction for given separation distance. The first stage is the long-range double layer interaction between the negatively charged silicon nitride AFM tip and the negatively charged cell surface, resulting in a repulsive force that decays exponentially away from the surface. This regime of the force profile can be fitted to double layer theory using the algorithm of Chan et al.

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to yield the Debye length of the

system. As stated, this regime of the force profile was small in the present study since the concentration of electrolyte is 10 mM with an expected Debye length of 3 nm, so can be safely neglected.


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Figure 2. Typical force profiles (red) of K. pneumoniae cells in: (a) HEPES buffer, and (b) peptide solution (90 minutes exposure time) with Pincus (black), Hertz (green) and Hooke’s law fit (blue) and derived parameters (capsule thickness, turgor pressure, polymer brush length and Young’s modulus) shown. Insets display data with fits on a normal axis.

Once the AFM tip makes contact with the bacterial surface, the tip indents the capsular polysaccharides. This region of the force curve can be fitted to the Pincus model

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for the

compression of a polyelectrolyte brush grafted to a rigid surface:


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= ln

(1)

where is the load applied to compress the surface, is the onset of linear compliance, and is a numerical prefactor, given by:

∝

(2)

where is the temperature, d is the grafted interchain distance,

!

is the Boltzmann constant,

and "! is the number of monomers carrying an ionic charge.

Under the experimental conditions used here, the temperature was constant within ±2% C, so that the magnitude of is determined by the ratio of "! ⁄'( . is, therefore, inversely proportional to the capsule density. If the region of the force curve corresponding to the compression of the capsule polymer brush is well fit by the Pincus model, the thickness of the capsule that is organised as a polymer brush can be determined. The distance between the onset of capsule indentation and the linear compliance region of the force profile is a reasonable estimate of the total capsule thickness.

The next stage of indentation is well described by the Hertz model.

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The Hertz model

describes the non-linear, elastic compression of a soft material (i.e., the cell wall and capsule polymers) by an infinitely hard, conical indenter (i.e., the AFM tip), viz: *+ *

-

89

= , ./ 12 23 tan 67

(3)

where is the loading force, ν is the Poisson ratio, : is Young’s modulus, and α is the cone half-opening angle of the AFM tip. A value of ν= 0.5 was used here to reflect an elastic sample 49 and α= 25º was specified by the manufacturer (Bruker, Camarillo, CA).


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The final stage of indentation corresponds to the point at which the loading force on the cell is high enough and indentation is that the turgor pressure is the dominant opposing force. This region of the force profile is very well fitted by Hooke’s law: 49, 55 = ;?@AB

(4)

where is the applied loading force, ;?@AB is the bacterial spring constant, and δ is the indentation. The slope of this linear compliance region of the force profile yields ;?@AB , which is a direct measure of the relative turgor pressure of the bacterium.

Figure 2(b) shows a typical force profile for the indentation of a K. pneumoniae cell after exposure to peptide (1.2 µM) for 83 minutes. Note that the form of the force profile is similar to that seen prior to the addition of peptide (Figure 2(a)) but the parameters obtained from the fits to the difference regimes of nanoindention are different. These changes report on how the peptide affects the capsule, cell wall and cytoplasmic turgor, and have been measured in time lapse for approximately 130 minutes exposure to peptide. These data for a typical cell plus AFM images of cells after long time peptide exposure are shown in Figures 3, 4 and 5 and are discussed below.

Figure 3 contains plots of the changes in turgor pressure and cell wall stiffness with time of exposure to peptide for a typical representative bacterial cell. After the first 42 minutes of peptide exposure, a turgor pressure of 300 mN/m was measured, which is significantly higher than the average turgor pressure of cells in buffer (98 mN/m). Concomitant with this is a dramatic increase in the cell wall and compressed capsule stiffness, reflected by a Young’s modulus of 15 kPa, which is significantly higher than the average Young’s modulus of cells in buffer (3.3 kPa). It might be that the increase in cell wall stiffness is simply due to the higher cytoplasmic turgor, increasing the internal pressure on the cell wall. However, at


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approximately 40 minutes exposure, the cell wall stiffness decreased and then stablizes over a period of around 60 minutes at ~10 kPa, while the turgor pressure continued to increase to a maximum of 474 mN/m at 116 minutes exposure time. The reduction in turgor pressure after this point suggests that lysis has occurred and cytoplasm has leaked from the compromised envelope/membrane. This observation is in good agreement with those of Liu et al.56 who observed a reduction in measured turgor pressure after exposing both Escherichia coli and Bacillus subtilis to antibacterial carbon nanotubes.

The changes in turgor pressure demonstrated by this data strongly suggest that the peptide is accumulating at the membrane. If the peptide were accumulating within the capsule or LPS, a change in the linear region of the force curve that reports on turgor pressure would likely not be evident. The initial increase in Young’s modulus supports this, as it implies that a change in the cell envelope structure has occurred, perhaps in terms of membrane fluidity or even peptidoglycan structure. Recent work by Saenz et al.

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shows that bacterial cell membranes

are organised in liquid ordered states by the presence of diplopterol, a bacterial ‘sterol surrogate’. The presence of amphipathic peptides could disrupt this organisation leading to a change in membrane fluidity.


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Figure 3. AFM data from force measurements on a single K. pneumoniae cell (representative data for one of five single cell experiments demonstrating the same trends): (a) a gradual increase in turgor pressure with a maximum at ~110 minutes and then a gradual decrease; and (b) an initial increase in Young’s modulus with little change from ~50 minutes until after ~110 minutes exposure time, where the Young’s modulus decreased. Error bars represent one standard deviation from the mean. Force data shown at time zero (i.e., before the addition of peptide) were acquired by measuring 10 force curves per cell across a population of cells and averaging the values obtained.

At approximately 110 minutes after peptide exposure, both turgor pressure and cell wall stiffness began decreasing at around the same rate. This corresponds to a dramatic change in


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the cell morphology as shown in the AFM images of Figure 4, which shows that the cells became somewhat bloated, misshapen and lose structural integrity, indicating possible cell death. This is supported by fluorescence viability assays that indicated that only ~10% of bacterial cells remained viable after 2 hours exposure to the peptide (see Supporting Information, Figure S2). Some variation in the kinetics of loss of turgor pressure and other characteristics was observed; this is not surprising and is an expectation for quantitative measurements of living systems since there is natural variation within the population due to e.g. the bacterial growth phase of individual bacteria within bacterial populations37, 58.

a

b

Figure 4. 20 × 20 µ images of K. pneumoniae cells submerged in: (a) HEPES buffer, and (b) 1.2 µM peptide solution after 100 minutes of exposure.

This study generated data in support of the hypothesis that the polymer brush length and capsule thickness did not change for the first 115 minutes exposure to the peptide solution (Figure 5). A change in capsule organisation was evident only after 115 minutes, corresponding to the rapid reduction in both and cell stiffness and turgor pressure observed between 115 and 130 minutes (Fig. 3). These data and the morphological changes that can be observed when imaging cells at high exposure times (Fig. 4) suggest that at these times cell death had occurred. This is an interesting observation for encapsulated bacteria because the


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capsule is held as an important part of the bacterial cell’s defense. The result is surprising since a cationic peptide should associate electrostatically with a putatively anionic polysaccharide capsule.59 A reduction in polysaccharide segment-segment repulsion and a resulting collapse of the capsule thickness and brush-like structure, therefore, would be expected. Such changes were not observed. We did, however, observe significant change in cytoplasmic turgor pressure and cell wall stiffness over the course of the experiment suggesting that the peptide interacts with the cells by rapidly translocating through the capsule, but clearly without any significant or long-lived electrostatic association. These observations suggest that even though the long-range electrostatic attraction between the peptide and capsule may first attract the peptide to the bacterial cell, it is the entropically driven association with the membrane that dominates, once the peptide approaches the bacterial outer envelope.


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Figure 5. AFM data collected from force measurements performed on a single K. pneumoniae cell (representative data for one of five single cell experiments demonstrating the same trends): (a) shows no significant change in polymer brush length; and (b) shows no significant change in capsule thickness until after ~120 minutes exposure time. Change after ~120 minutes corresponds to the rapid change shown in Fig. 3 that suggests cell death has occurred.

To further investigate the role of the capsule in mediating cell-peptide interactions, imaging and force measurements were conducted on capsule-deficient K. pneumoniae cells, for direct comparison (Figure 6). The morphology of capsule-deficient cells was comparable to that of


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the wild type cells (data not shown). Force curves obtained were highly reproducible and the difference between the capsule deficient and wild-type cells only became apparent when taking force measurements: the force curves of the easily compressible wild-type cells and the rigid capsule deficient cells are shown for comparison in Figure 6(a).

Figure 6: (a) Typical force profile of a capsule-deficient K. pneumoniae cell and a wild type K. pneumoniae cell in 1.2 µM peptide solution (~90 minutes exposure time) with Hooke’s law fit and derived parameters; and (b) Turgor pressure of capsule deficient K. pneumoniae cells. Force data shown at time zero (i.e., before the addition of peptide) were acquired by measuring 10 force curves per cell and averaging the values obtained. Note the measured capsule thickness of 5 nm for the capsule deficient cell in (a) most likely reflects the presence


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of other polymer-like structure on the cell surface, such as lipopolysaccharides, rather than the polysaccharide that makes up the bulk of the capsule.

Fitting the force profiles for the capsule-deficient mutants to Hooke’s Law to obtain a measure of the cytoplasmic turgor, as reported by Wang et al., 32 showed that cells without a capsule were under extreme osmotic stress as indicated by the turgor pressure of 300 mN/m for the cell prior to exposure to peptide (Fig. 6). After exposure to the peptide, there was a continual reduction in turgor pressure for the duration of peptide exposure, i.e., 120 mins. The capsule-deficient cells did not adhere to the gelatin-coated surface with sufficient avidity to withstand multiple force profile collections. Therefore, force curves of capsule-deficient cells were collected across a population of cells. As such, direct comparisons between the single cell data shown in Figures 3 and 5 and the capsule deficient population of cells shown in Figure 6(b) cannot be made due to the differing experimental methodology. That said, the data in Figure 6(b) demonstrate that the peptide compromised the capsule-deficient bacterial envelopes. Fluorescence live/dead cell assays were more difficult to perform on the capsuledeficient cells since loss of capsule reduces adhesion to coated slides and the live/dead assay methodology requires several washing stages. However, the studies showed that the cell membranes were compromised on exposure to peptide to a similar extent in the presence or absence of capsule. For comparison, the MIC and MBC for the peptide was measured for both wild type and capsule-deficient bacteria and found to be 38µM in all cases. That the MIC and MBC are equal confirms that the bacterial cells are dying as a result of the peptide induced membrane damage. That the MIC and MBC of the capsule-deficient and wild-type cells are equal is further evidence that the capsule does not play a large role in the collapse of the cell due to cytoplasm leakage. Unlike the case for some K. pneumoniae isolates studied


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by Campos et al.60, the capsule does not appear to be a significant factor for determining susceptibility to our model AMP for the K. pneumoniae isolate AJ218.

Conclusions

The turgor pressure of K. pneumoniae cells increased upon addition of the melittin (P14K) peptide, likely due to its accumulation at the cytoplasmic membrane causing lysis. This increase in turgor pressure was accompanied by an increase in Young’s modulus, which suggests a stiffening of the cell envelope due to accumulation of peptide. The data presented here show that K. pneumoniae AJ218 capsule was largely unaffected by exposure to the peptide, which indicates that ionic interaction of bacteria-bound capsular polysaccharide is not a key factor for AMP interaction with K. pneumoniae AJ218 cells. The capsule, however, does not prevent cell lysis as seen in the case of the mutant, which had a similar response to the peptide. The K54 capsule does not appear to offer protection to Klebsiella pneumoniae AJ218 against antimicrobial peptides.

Acknowledgements

The authors gratefully acknowledge the support of the Australian Research Council and the National Health and Medical Research Council including the NHMRC Program Grant in Cellular Microbiology. Research at The Florey Institute of Neuroscience and Mental Health is supported by the Victorian Government Operational Infrastructure Support Program. AM received an Australian Postgraduate Award. All AFM work was performed in part at the Materials Characterization and Fabrication Platform (MCFP) at the University of Melbourne.


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Supporting Information Available

A schematic representation of the mechanical model for cell indentation utilised in this work, developed by Wang et al.32 appears in Figure S1. The results of cell viability assays, performed with wild-type K. pneumoniae AJ218 in HEPES buffer and peptide solution, appear in Figure S2. Force curves measured on both wild-type and capsule deficient K. pneumoniae AJ218 cells in buffer and peptide solution are overlaid in Figure S3 to demonstrate the reproducibility of force curves collected. This information is available free of charge via the Internet at http://pubs.acs.org.

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Atomic Force Microscopy Reveals the Mechanobiology of Lytic

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