Adhesion Mechanism of the Staphylococcal Polysaccharide

Sticky Matrix: Adhesion Mechanism of the Staphylococcal Polysaccharide Intercellular Adhesin Cécile Formosa-Dague,†,¶ Cécile Feuillie,†,¶ Audrey Beaussart,† Sylvie Derclaye,† Soňa Kucharíková,‡,§ Iñigo Lasa,∥ Patrick Van Dijck,‡,§ and Yves F. Dufrêne*,†,⊥ †

Institute of Life Sciences, Université catholique de Louvain, Croix du Sud, 4-5, bte L7.07.06., B-1348 Louvain-la-Neuve, Belgium Department of Molecular Microbiology, VIB, and §Laboratory of Molecular Cell Biology, Institute of Botany and Microbiology, KU Leuven, 3000 Leuven, Belgium ∥ Group of Microbial Communities and Disease, Navarrabiomed-FMS, UPNA, IdiSNA, 31008 Navarra, Spain ⊥ Walloon Excellence in Life sciences and Biotechnology (WELBIO), 1300 Wavre, Belgium ‡

ABSTRACT: The development of bacterial biofilms on surfaces leads to hospital-acquired infections that are difficult to fight. In Staphylococci, the cationic polysaccharide intercellular adhesin (PIA) forms an extracellular matrix that connects the cells together during biofilm formation, but the molecular forces involved are unknown. Here, we use advanced force nanoscopy techniques to unravel the mechanism of PIA-mediated adhesion in a clinically relevant methicillin-resistant Staphylococcus aureus (MRSA) strain. Nanoscale multiparametric imaging of the structure, adhesion, and elasticity of bacteria expressing PIA shows that the cells are surrounded by a soft and adhesive matrix of extracellular polymers. Cell surface softness and adhesion are dramatically reduced in mutant cells deficient for the synthesis of PIA or under unfavorable growth conditions. Single-cell force spectroscopy demonstrates that PIA promotes cell−cell adhesion via the multivalent electrostatic interaction with polyanionic teichoic acids on the S. aureus cell surface. This binding mechanism rationalizes, at the nanoscale, the well-known ability of PIA to strengthen intercellular adhesion in staphylococcal biofilms. Force nanoscopy offers promising prospects for understanding the fundamental forces in antibiotic-resistant biofilms and for designing anti-adhesion compounds targeting matrix polymers. KEYWORDS: Staphylococcus aureus, polysaccharide intercellular adhesin, adhesion forces, biofilm matrix, atomic force microscopy

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poly-N-acetylglucosamine (PNAG), plays a key role in cell− cell adhesion. 9−11 Positive charges introduced by the deacetylation of N-acetylglucosamine residues are believed to bind to negatively charged cell surface molecules. Currently, two unsolved questions are what is the ligand of PIA, and what are the molecular forces involved? The presence of PIA is not essential for biofilm development since several S. aureus isolates are able to produce protein matrices.12,13 Interestingly, S. aureus strains have been shown to regulate the production of different types of matrices depending on environmental conditions, thus enabling them to readily switch between polysaccharide and protein-based biofilms.14 Atomic force microscopy (AFM) has offered new opportunities to understand the forces involved in cell adhesion and biofilm formation.15,16 Multiparametric imaging has enabled the surface structure and properties of live cells to be probed,

he formation of microbial biofilms on surfaces is of major relevance to medicine and industry.1,2 Bacterial cells in a biofilm produce a matrix of extracellular polymeric substances containing polysaccharides, proteins, nucleic acids, and lipids that attaches the cells to surfaces, holds them together, and ensures the mechanical stability of the biofilm.3−6 Among these polymers, extracellular polysaccharides represent a major component of biofilm matrices. Despite the key roles of biofilm matrix polymers, how they connect the cells together to form biofilms is largely unknown at the molecular level.5 A better knowledge of the fundamental forces in biofilm matrices would increase our understanding of the cohesive strength and stability of biofilms. The development of biofilms by Staphylococcus aureus is a major cause of nosocomial infections. Following initial adhesion of the bacteria to host cells and protein-coated materials, cell− cell adhesion (aggregation) and multiplication lead to the formation of a mature biofilm in which the cells are surrounded by extracellular matrix polymers. 7,8 Among these, the polycationic polysaccharide intercellular adhesin (PIA), or © XXXX American Chemical Society

Received: November 28, 2015 Accepted: February 18, 2016

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DOI: 10.1021/acsnano.5b07515 ACS Nano XXXX, XXX, XXX−XXX

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ACS Nano including bacteria.17−21 In this newly developed modality, spatially resolved force curves are recorded at high frequency to generate adhesion and mechanical maps of the cell surface with high spatiotemporal resolution. In single-cell force spectroscopy (SCFS), the AFM tip is replaced by a living cell in order to measure single-cell adhesion forces.22 A novel colloidal probe assay has recently enabled the reliable analysis of bacterial cell adhesion forces.23,24 A colloidal particle is attached to the end of a cantilever and coated with a bioinspired polydopamine wet adhesive. The sticky colloidal probe is used to attach a single live cell, then force−distance curves are acquired on a target surface, such as a solid substrate or another cell. This approach has provided novel insights into the forces involved in the adhesion of a variety of medically important microbes.15,16 Here, we use these modalities to explore the nanoscale forces driving PIA-dependent adhesion. We focus on the clinical methicillin-resistant S. aureus (MRSA) strain 132, which is able to switch between PIA and proteinaceous biofilm matrices depending on environmental conditions.14 We show that, under favorable growth conditions, intercellular adhesion originates from electrostatic interactions between PIA and teichoic acids on the S. aureus cell surface.

Figure 2. PIA is involved in biofilm formation. (a) Crystal violet staining of biofilms made of WTPIA and ΔPIA cells grown in PIA conditions. ΔPIA cells showed significantly reduced biofilm biomass in comparison with the WT (*p < 0.05). (b) Fluorescence images after staining with FITC-WGA, documenting the presence of a PIA matrix (green) in WTPIA cells, both in the biofilm (main image) and in the planktonic (bottom left inset) stages. (c) Biofilms of ΔPIA cells were hardly labeled. (d) Similarly, no fluorescence was detected for biofilms of WTFnBP cells grown in the presence of 0.2% glucose (FnBP conditions). The top insets in (bd) are DIC images. Scale bars: 10 μm.

RESULTS AND DISCUSSION Role of PIA in Cell Adhesion and Biofilm Formation. We initially confirmed the involvement of PIA in S. aureus adhesion and biofilm formation using microscopic assays. Figure 1a shows that S. aureus 132 cells grown in conditions

agglutinin, specific for N-acetyl-D-glucosamine, and conjugated to fluorescein isothiocyanate (FITC), and stained cells were then imaged using confocal microscopy (Figure 2b−d). Biofilms of WTPIA cells (i.e., biofilms formed under 3% NaCl conditions) featured strong fluorescence (Figure 2b), as opposed to biofilms of ΔPIA cells (Figure 2c). Fluorescence was also observed between WTPIA cells, indicating that PIA formed a matrix connecting the bacteria together. Interestingly, planktonic WTPIA cells were also labeled (Figure 2b, bottom left inset), meaning that these cells produced PIA both in biofilm and planktonic conditions. Finally, fluorescence was hardly detected for biofilms made of WT cells grown in 0.2% glucose conditions, which favor the production of fibronectinbinding proteins (FnBPs, WTFnBP cells; Figure 2d), a finding consistent with the ability of this strain to switch between PIA and proteinaceous biofilm matrices.14 These observations confirm that, under appropriate environmental conditions, MRSA strain 132 produces a matrix of PIA that promotes intercellular adhesion during biofilm formation. It has been suggested that the positive charges of PIA may bind to negatively charged cell surface polymers such as teichoic acids (Figure 1, right cartoons), but direct evidence for such interaction is lacking. Therefore, we sought to investigate the molecular mechanism (binding strength, binding specificity) behind PIA-dependent adhesion. Nanoscale Structure, Elasticity, and Adhesion of PIA. We explored the nanoscale structural and biophysical properties of the PIA matrix directly on living bacteria, using multiparametric imaging. Shown in Figure 3 are correlated images of the structure, elasticity, and adhesion of S. aureus WTPIA cells. Unless stated otherwise, bacteria were not centrifuged in order

Figure 1. Role of PIA in intercellular adhesion. (a,b) Optical microscopy images of S. aureus WTPIA cells (a) and ΔPIA cells (b), cultured in PIA conditions (trypticase soy broth (TSB) + 3% NaCl) and resuspended in 100 mM NaCl. As illustrated in the schematics (right panels), the formation of large aggregates by WTPIA cells is thought to involve the electrostatic interaction between cationic PIA (pink) and negatively charged molecules on the S. aureus cell surface (blue).

favoring the production of PIA (3% NaCl, hereafter “WTPIA cells”) formed large aggregates. By contrast, cells from a 132 ΔicaADBC deletion mutant lacking PIA (“ΔPIA cells”) grown in the same conditions were isolated, without any detectable aggregates (Figure 1b). We also assessed the ability of these strains to develop biofilms on polystyrene plates under PIA conditions (3% NaCl), using crystal violet staining. Compared to WTPIA cells, ΔPIA cells showed significantly decreased biofilm biomass (Figure 2a; *p < 0.05). To confirm that PIA was properly expressed at the cell surface, bacteria were labeled with the lectin wheat germ B


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electrostatic interactions between the positively charged PIA matrix and negative charges on the silicon nitride tip. To provide direct evidence that the observed structural and physical properties are associated with PIA, correlated images of the structure, elasticity, and adhesion were recorded for S. aureus ΔPIA cells lacking PIA and for S. aureus WTFnBP cells grown in conditions favoring the production of FnBPs. As can be seen in Figure 5, the cells were no longer surrounded by a thick extracellular layer (Figure 5a−d) and showed stiff properties (Figure 5e−h). The Young’s modulus obtained on top of WTFnBP cells and ΔPIA cells was 538 ± 266 and 542 ± 303 kPa (Figure 4c−f), thus similar to that of centrifuged WTPIA cells. We note that softer elasticity was observed on the edges of the cells, an artifact associated with the poorly controlled contact geometry (Figure 5e−h). While ΔPIA cells hardly adhered to the negatively charged tip (Figure 5k,l), there was substantial adhesion on WTFnBP cells (Figure 5i,j), a behavior that may reflect binding of FnBPs to the tip. In a nutshell, these data show that, under appropriate growth conditions, S. aureus strain 132 produces copious amounts of extracellular PIA that dramatically increases the softness and stickiness of the cell surface. Strength and Specificity of PIA-Mediated Intercellular Adhesion. To gain insight into the adhesion mechanism of PIA, we quantified the forces between individual bacteria using SCFS (Figure 6). In Figure 6a,b, we present the maximum adhesion forces, rupture distances, and representative force signatures recorded in 100 mM NaCl at short contact time (100 ms) for three pairs of S. aureus WTPIA cells. The maximum adhesion force corresponds to the largest adhesion event in each force curve, while the rupture length is the distance of the last adhesion event. Many retraction force profiles (79%) showed multiple binding events with a maximum adhesion force of 148 ± 38 pN and a rupture length of 143 ± 31 nm (n = 4000 curves from 10 cells; Table 1). The extended ruptures and the lack of defined force patterns are consistent with the stretching of long polysaccharides. Multiple adhesion peaks may be due to the interaction of either multiple loops of the same polymer or multiple PIA molecules. The approach curves did not show any particular attractive or repulsive feature. Adhesion was abrogated between ΔPIA cells (Figure 6g,h), confirming that the forces measured between WTPIA cells are mediated by PIA. Different WTPIA cells from independent cultures showed similar adhesion force profiles (Table 1), suggesting that the cell population was homogeneous. Yet it is important to note that substantial variations in adhesion properties were observed when using cells cultured from agar plates that were older than 2 weeks. Instability in PIA production is known to occur in some strains of S. aureus, very likely because the synthesis of PIA is energy-consuming and PIA-negative variants are easily selected. This phenotype instability has been associated with the insertion of IS256 in the icaADBC operon.25 We observed an increase in adhesion frequency, adhesion force, and rupture length when increasing the contact time to 1 s (Figure 6c,d and Table 1). In line with earlier studies on staphylococcal adhesion,26−29 this effect is likely to originate from an increased number of molecular bonds with interaction time. In the PIA context, we suggest that the time dependency reflects the time necessary to achieve optimal fitting between multiple positive charges of PIA chains and negatively charged molecules on opposing cells. The longer extensions are

Figure 3. Multiparametric imaging of the structural and biophysical properties of the PIA matrix. (a,b) Height images (color scale: 2 μm) of S. aureus WTPIA cells cultured in TSB 3% NaCl and recorded in Tris-buffered saline (50 mM Tris, 150 mM NaCl, pH 7.4), and simultaneous (c,d) elasticity images (color scale: arbitrary units) and (e,f) adhesion images (color scale: 500 pN). Scale bars: 1 μm. Labels “M” and “C” refer to the PIA matrix and to the cell surface, respectively. The dotted line indicates a change in image contrast, resulting from strong tip−PIA interactions.

to prevent removal of PIA molecules from the cell surface. Topographic images revealed that the cells (Figure 3a, label C) were surrounded by a fuzzy layer of extracellular polymers (Figure 3a, label M), about twice as large as the cell size (∼2 μm in diameter). In light of control experiments (see below), we attribute this layer to the PIA matrix. For some cells, abrupt changes in topographic contrast were observed (Figure 3a, dotted line), indicating that the tip had pushed the extracellular layer away and contacted the rigid cell wall. Consistent with this, the extracellular layer featured a very low elasticity, except in localized regions were the tip was in direct contact with the stiff cell wall (Figure 3c,d). To obtain quantitative information on the cell surface elasticity, force versus indentation curves were recorded on native WTPIA cells, and on WTPIA cells from which the PIA layer had been removed by centrifugation (Figure 4a,b). The Young’s modulus was assessed by fitting the curves using the Hertz model, yielding values of 45 ± 27 and 520 ± 173 kPa (mean ± SD on a total of n = 5120 curves from five different cells from two cultures) for native and centrifuged cells, respectively. This result demonstrates that PIA forms a soft extracellular layer around the cells. Adhesion images were highly contrasted, in that adhesion events of 122 ± 10 pN (mean ± SD on a total of n = 9313 curves from three different cells) were mostly detected on the soft extracellular layer (see bright pixels in Figure 3e, label M, and Figure 3f), not on the cell wall. So PIA conveys soft and sticky properties to the cell surface. We speculate that the measured adhesion results from C


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Figure 4. Nanomechanical analysis shows that PIA forms a very soft layer surrounding the cells. (a,c,e) Representative force−indentation curves and (b,d,f) histograms of Young’s modulus values obtained on 500 nm × 500 nm areas on S. aureus WTPIA cells in native conditions or after centrifugation (a,b), on native WTFnBP cells (c,d), and on native ΔPIA cells (e,f), recorded in 100 mM NaCl. Shown in red are the fits obtained with the Hertz model on the first 50 nm of indentation. The blue line (in a) and blue histogram (in b) are for native S. aureus WTPIA cells. For each condition, data from a total of n = 5120 curves from five different cells are shown.

consistent with a model in which, with time, longer fractions of the PIA chains become engaged in intercellular bonds. As PIA is thought to bind through electrostatic interactions, we hypothesized that cell−cell adhesion forces should vary with ionic strength. Increasing the concentration of monovalent salts to 500 mM indeed abolished the adhesion between WTPIA cells (Figure 6e,f). Consistent with the behavior of polyelectrolytes, high ionic strengths are expected to lead to collapsed conformations of the PIA chains in which the charges are shielded, thus decreasing the probability to form intercellular bonds. The dependence of PIA adhesion on environmental conditions could be of biological relevance, providing a means to modulate the stability and dispersal of PIA-dependent biofilms. We also found that the measured intercellular forces dramatically depend on growth conditions. Figure 7a,b shows that the adhesion between WTPIA and WTFnBP cells (307 ± 79 pN, n = 2000 curves from five cells) was stronger than that

between two WTPIA cells. Presumably, the positively charged PIA molecules on WTPIA cells will bind more strongly to PIAdeficient cells than to PIA-rich cells, where electrostatic repulsion between the two PIA layers may occur. By contrast, no adhesion was observed between WTFnBP cells (Figure 7c,d; adhesion frequency of ∼3% for five different cell pairs), which is somewhat surprising as FnBP proteins are known to mediate intercellular adhesion.30,31 This suggests that, under our analysis conditions, FnBPs are not fully functional or that their adhesive properties cannot be captured by AFM. What is the molecular basis of PIA-based adhesion? It has been proposed that PIA connects the cells together by electrostatic interaction between its positively charged groups and negatively charged molecules on opposing cells. As wall teichoic acids (WTAs) are the most abundant polyanions of the cell wall and play an important role in S. aureus adhesion,32,33 they have been suggested to serve as an important ligand for PIA-mediated adhesion.7 To demonstrate a direct interaction D


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Figure 5. Cells lacking PIA show major differences in nanobiophysical properties. (a−d) Height images (color scale: 2 μm) and corresponding (e−h) elasticity images (color scale: arbitrary units) and (i−l) adhesion images (color scale: 500 pN) in Tris-buffered saline of S. aureus WTFnBP cells cultured in TSB 0.2% glucose and of S. aureus ΔPIA cells cultured in TSB 3% NaCl. Scale bars: 1 μm.

It is intriguing to understand how electrostatic repulsion between PIA-positive bacteria is overcome in nature. One possibility is the existence of heterogeneity in the amount of PIA produced by the bacterial population. Thus, PIA-producing bacteria interact with those bacteria that produce low levels of PIA. It is also possible that bacterial interactions occur before large amounts of PIA surround the cell wall or that PIA does not cover the cell homogeneously, so that interactions would take place between PIA-covered and PIA-naked regions of neighbor cells. Deacetylation may be of biological relevance in other organisms. For instance, Pga production in Escherichia coli also requires the participation of PgaB, a deacetylase homologous to IcaB. In the case of fungal pathogens of plants, deacetylated chitin (chitosan) from the microbial cell wall induces the rapid influx of calcium and the rapid induction of cell wall modifications and expression of PRP proteins.38 It is believed that there might be a specific receptor for short glucosamine fragments.

between PIA and WTAs, we measured the forces between PIAproducing cells and mutant cells deficient for the production of WTAs (“ΔWTA cells”). WTA depletion was achieved in S. aureus through the inactivation of the tagO gene that encodes the first enzyme in the WTA biosynthesis pathway.34−36 As can be seen in Figure 8a,b, adhesion forces were abrogated between WTPIA and ΔWTA cells, thus demonstrating that WTAs represent a major ligand of PIA. This finding explains why WTAs often contaminate PIA samples during purification.36,37 However, why PIA from ΔWTA cells did not bind to WTAs from WTPIA cells? One explanation is that PIA in the WTA mutant may not be properly attached to the cell wall and, in this state, is unable to promote efficient interactions with the WT. To provide further evidence for the electrostatic nature of the PIA interaction, we measured the forces between WTPIA cells and negatively charged model substrates (Figure 8c−f). All the curves recorded in 100 mM NaCl on negatively charged carboxyl-terminated surfaces featured strong adhesion forces, with multiple force peaks of 1185 ± 144 pN magnitude and 193 ± 28 nm rupture lengths (Figure 8c,d; n = 2000 curves from five cells). The extended rupture lengths, similar to those measured between two cells, indicate that long polysaccharides were stretched. Much weaker adhesion (228 ± 21 pN) and shorter extensions (113 ± 42 nm) were observed when increasing the salt concentration (Figure 8e,f) or when using neutral hydroxyl surfaces (Figure 8g,h), thus strongly supporting the notion that cationic PIA chains bind to anionic surfaces via multivalent electrostatic interactions. As most natural and biomaterial surfaces are negatively charged, this finding suggests that PIA favors cell−substrate interactions. In summary, these results provide direct evidence that, during biofilm formation, PIA mediates cell−cell association and cell− substrate adhesion via electrostatic interactions.

CONCLUSIONS During the development of biofilms, staphylococcal species produce extracellular matrix polymers (exopolysaccharides, proteins, and eDNA) that surround and connect the cells together.8,13 While PIA represents the most widely investigated matrix component, how this macromolecule holds bacteria together to form a stable biofilm is poorly understood. It is therefore of great importance to shed light into the molecular basis of PIA-mediated adhesion. Using nanoscopy tools, we have measured the forces in PIA-dependent adhesion at the single-cell level, thereby providing compelling evidence that this polymer binds WTAs on the S. aureus cell surface through multivalent electrostatic forces. Under appropriate growth conditions (NaCl), S. aureus produces an extracellular layer of PIA that dramatically increases the softness and adhesiveness of E


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Figure 6. Forces driving PIA-dependent cell−cell adhesion. (a,c) Adhesion force and (b,d) rupture distance histograms obtained at 0.1 s (a,b) or 1.0 s (c,d) contact time in 100 mM NaCl for three pairs (three different colors) of S. aureus WTPIA cells. (e,f) Force data obtained at 0.1 s contact time for three pairs of S. aureus WTPIA cells in 500 mM NaCl. (g,h) Force data obtained at 0.1 s contact time for three pairs of S. aureus ΔPIA cells in 100 mM NaCl. The insets in (b,d,f,h) show representative force signatures; “n.a.” values are the percentages of nonadhesive events. All curves were obtained using an applied force of 250 pN and an approach and retraction speed of 1.0 μm/s. The cartoons highlight key surface molecules, i.e., positively charged PIA (pink) and negatively charged wall teichoic acids (blue).

the cell surface. PIA mediates time-dependent intercellular forces that originate from electrostatic interactions between cationic PIA polyelectrolytes and anionic WTAs on neighboring cells. Increasing the ionic strength leads to weaker cell−cell adhesion, revealing that at high salt concentration PIA chains are collapsed and lose their polyelectrolyte properties. We speculate that the multivalent and polyelectrolyte nature of

PIA−WTA bonds plays a role in increasing the lifetime and stability of cellular contacts, thus enabling S. aureus to form cohesive and stable biofilms. PIA also promotes electrostatic interactions with negatively charged substrates. Change in growth conditions (NaCl- vs glucose-rich conditions) profoundly alters the structural, mechanical, and adhesive phenotype of the S. aureus cell surface. Our data emphasize F


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matrix. For instance, compounds capable of modulating electrostatic forces in PIA matrices could be used to inhibit staphylococcal biofilm formation.

Table 1. Probability of Adhesion (Padh = % of curves with adhesion events), Adhesion Force (Fadh), and Rupture Length (Ladh) Measured for Different Pairs of S. aureus WTPIA Cells Using an Interaction Time of 0.1 or 1.0 s 0.1 s

pair 1 pair 2 pair 3 pair 4 pair 5 pair 6 pair 7 pair 8 pair 9 pair 10 mean SD

METHODS

1.0 s

Padh (%)

Fadh (pN)

Ladh (nm)

Padh (%)

Fadh (pN)

Ladh (nm)

77.0 73.9 62.9 86.0 53.5 78.3 89.0 91.8 78.6 99.3 79.0 13.6

128.4 159.0 170.1 187.1 111.9 110.3 160.8 125.8 103.7 221.6 147.9 38.4

125.5 157.6 116.6 184.0 145.2 104.4 192.2 117.3 117.2 171.3 143.1 31.5

98.3 99.3 99.8 99.8 99.3

347.0 390.8 454.7 295.1 354.9

200.3 217.6 214.6 218.7 241.0

99.3 0.6

368.0 59.1

218.4 14.6

Strains and Culture Conditions. The following S. aureus strains were used in this study: S. aureus strain 132 (MRSA clinical strain), S. aureus strain 132 Δica (132 with deletion of the icaADBC operon),14 and strain 15981 ΔtagO (with deletion of the tagO gene).36 Strains 132 and 132 Δica were cultured in Trypticase soy broth (TSB) supplemented with either 0.2% glucose (FnBP condition) or 3% NaCl (PIA condition) for 12 h at 37 °C under agitation (180 rpm). The ΔtagO strain was cultured in the same conditions at 30 °C. Microscopic Adhesion Assay. Aggregation phenotypes were directly observed after resuspension of the cells in 5 mL of 100 mM NaCl. Aggregation levels were observed by optical microscopy (bright field) using an Axio Observer Z1 microscope with a 40× objective (Zeiss, Germany). Biofilm Development. For in vitro biofilm biomass determination, S. aureus cells were collected from TSB plates and dissolved in sterile water. Biofilms were studied in flat-bottom 96-well polystyrene plates (Greiner Bio-one, Germany). Briefly, cells were adjusted to a final concentration of 1 × 105 cells mL−1 in TSB supplemented with 3% NaCl (PIA condition) and allowed to form mature biofilms on the polystyrene substrate (24 h, 37 °C, static). Afterward, non-biofilmassociated cells were removed by washing with phosphate-buffered saline (PBS) and processed for crystal violet staining as described earlier.39 Remaining biofilms were air-dried for approximately 10 min and subsequently stained with crystal violet (1% solution prepared in sterile water, 200 μL/well) for 15 min at room temperature. The remaining solution was removed by three rounds of washing with sterile water (200 μL/well). Then, wells were air-dried for 10 min.

the key role of the deacetylation of the N-acetylglucosamine residues of PIA: by introducing positive charges into the polymer, multiple interactions with negative charges on opposing cells are favored. Our nanoscale platform offers promising prospects for understanding the contribution of polysaccharide and protein matrices to biofilm formation in Staphylococci, including MRSA strains, and for developing novel drugs targeting the

Figure 7. Growth conditions profoundly influence intercellular adhesion. (a) Adhesion force and (b) rupture distance histograms obtained in 100 mM NaCl at 0.1 s contact time between S. aureus WTPIA cells (grown in TSB 3% NaCl) and S. aureus WTFnBP cells (cultured in TSB 0.2% glucose). (c,d) Force data obtained in the same conditions for three pairs of S. aureus WTFnBP cells. The cartoons highlight key surface molecules, i.e., PIA (pink), wall teichoic acids (blue), and FnBPs (red). G


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Figure 8. Adhesion mechanism: PIA binds to teichoic acids via multivalent electrostatic forces. (a,b) Adhesion force and rupture distance histograms obtained in 100 mM NaCl between S. aureus WTPIA cells and S. aureus ΔWTA cells grown in TSB 3% NaCl. (c−f) Force data obtained in 500 mM NaCl (c,d) or 500 mM (e,f) between three S. aureus WTPIA cells and carboxyl-terminated substrates. (g,h) Force data obtained in 100 mM NaCl between three S. aureus WTPIA cells and hydroxyl-terminated substrates. All curves were obtained using a contact time of 0.1 s, an applied force of 250 pN, and an approach and retraction speed of 1.0 μm/s. Multiparametric Imaging. Cells were immobilized on polystyrene substrates. Images were recorded in Tris-buffered saline (TBS) using the Quantitative Imaging mode40 available on the Nanowizard III AFM (JPK Instruments, Germany), with MSCT cantilevers (Bruker, nominal spring constant of 0.01 N/m). Multiparametric images were recorded at 128 pixels × 128 pixels with an applied force kept at 0.65 nN for all conditions and a constant approach/retract speed of 90 μm/s (z-range of 2 μm). Young’s moduli were calculated using the Hertz model, in which the force (F), indentation (δ), and Young’s modulus (E) follow the equation F = (2 × E × tan α)/(π(1 − ν2)δ2), where α is the tip opening angle (17.5°) and ν the Poisson ratio (arbitrarily assumed to be 0.5).41 The cantilever spring constants were determined by the thermal noise method.42 For each condition, experiments were repeated for five cells from independent cultures. Single-Cell Force Spectroscopy. For cell−cell analyses, target cells were immobilized on polystyrene substrates. For cell−substrate analyses, carboxyl- and hydroxyl-terminated self-assembled monolayers (SAMs) were prepared by immersing gold-coated substrates in ethanol solutions containing 1 mM of 16-mercaptohexadecanoic acid (Sigma-

Residues of crystal violet were distained with acetic acid (33%, 180 μL/well). Biofilm biomass was measured using a spectrophotometer (Spectra max Plus 384) at 590 nm. Statistical analyses were performed using the Student’s t-test (GraphPad Prism software). Differences were considered significant if *p ≤ 0.05. All in vitro experiments performed in 96-well polystyrene plates were repeated twice, using six wells per strain. Confocal Scanning Laser Microscopy. For confocal microscopy, S. aureus biofilms were developed on plastic highly adhesive round tissue culture coverslips (13 mm diameter, Sarstedt, Germany). One milliliter of S. aureus suspension (1 × 105 cells) prepared in TSB supplemented with 0.25% glucose (FnBP condition) or 3% NaCl (PIA condition) was added to wells containing coverslips (one per well). Biofilms were incubated for 24 h at 37 °C. Mature biofilms were washed once with PBS and subsequently stained with FITC-WGA lectin (50 μg/mL) (GeneTex, USA) at 37 °C for 1 h (green fluorescence). Imaging was carried out with an Olympus FV1000 confocal laser scanning biological microscope, and images were processed with the FV10-ASW 2.0 software. H


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Aldrich, 98%) or 1 mM of 11-mercapto-1-undecanol (Sigma-Aldrich, 98%) overnight, rinsing them with ethanol, and drying them under N2. To prepare the cell probes, colloidal probes were obtained by attaching a single silica microsphere (6.1 μm diameter, Bangs Laboratories) with a thin layer of UV-curable glue (NOA 63, Norland Edmund Optics) on triangular tipless cantilevers (NP-O10, Bruker) and using a Nanowizard III AFM (JPK Instrument, Berlin, Germany). Cantilevers were then immersed for 1 h in TBS (pH 8.5) containing 4 mg/mL of dopamine hydrochloride (Sigma-Aldrich), rinsed in TBS, and used directly for cell probe preparation. The nominal spring constant of the colloidal probe cantilever was ∼0.06 N/m, as determined by the thermal noise method. Then, 50 μL of a diluted cell suspension was deposited into the Petri dish containing previously adhered cells or SAM substrates, at a distinct location within the Petri dish; 3 mL of NaCl (100 or 500 mM) was added to the system. The colloidal probe was brought into contact with an isolated bacterium and retracted to attach the bacterial cell. Proper attachment of the cell on the colloidal probe was checked using optical microscopy. Cell probes were used to measure cell−cell and cell−substrate interaction forces at room temperature, using an applied force of 0.25 nN, a constant approach− retraction speed of 1.0 μm/s, and a contact time of either 100 ms or 1 s. Data were analyzed using the Data Processing software from JPK Instruments (Berlin, Germany). Adhesion force and distance rupture histograms were obtained by calculating the maximum adhesion force and rupture distance of the last peak for each curve. Unless stated otherwise, for each condition, experiments were repeated for five cells from independent cultures.

AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]. Author Contributions ¶

C.F.-D. and C.F. contributed equally to this work.

Notes

The authors declare no competing financial interest.

ACKNOWLEDGMENTS Work at the Université catholique de Louvain was supported by the National Fund for Scientific Research (FNRS), the FNRSWELBIO under Grant No. WELBIO-CR-2015A-05, the Federal Office for Scientific, Technical and Cultural Affairs (Interuniversity Poles of Attraction Programme), and the Research Department of the Communauté française de Belgique (Concerted research action). Work at Navarrabiomed was supported by the Spanish Ministry of Economy and Competitiveness under Grant No. BIO2014-53530-R. Y.F.D. and C.F.-D. are, respectively, Research Director and Postdoctoral Researcher of the FRS-FNRS. S.K. was supported by an FWO postdoctoral grant. REFERENCES (1) Costerton, J. W.; Stewart, P. S.; Greenberg, E. P. Bacterial Biofilms: A Common Cause of Persistent Infections. Science 1999, 284, 1318−1322. (2) Kolter, R.; Greenberg, E. P. Microbial Sciences: The Superficial Life of Microbes. Nature 2006, 441, 300−302. (3) Xavier, J. B.; Foster, K. R. Cooperation and Conflict in Microbial Biofilms. Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 876−881. (4) Karatan, E.; Watnick, P. Signals, Regulatory Networks, and Materials That Build and Break Bacterial Biofilms. Microbiol. Mol. Biol. Rev. MMBR 2009, 73, 310−347. (5) Flemming, H.-C.; Wingender, J. The Biofilm Matrix. Nat. Rev. Microbiol. 2010, 8, 623−633. (6) Payne, D. E.; Boles, B. R. Emerging Interactions between Matrix Components during Biofilm Development. Curr. Genet. 2016, 62, 137−141. I


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Adhesion Mechanism of the Staphylococcal Polysaccharide

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