Differential Lipopolysaccharide Core Capping Leads to ...and architecture through modulation of cellular mechanics and microcolony structures, thereby contributing to bacterial com-munity

JOURNAL OF BACTERIOLOGY, Nov. 2009, p. 6618–6631 Vol. 191, No. 210021-9193/09/$12.00 doi:10.1128/JB.00698-09Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Differential Lipopolysaccharide Core Capping Leads to Quantitativeand Correlated Modifications of Mechanical and Structural

Properties in Pseudomonas aeruginosa Biofilms�†Peter C. Y. Lau,1,2 Theresa Lindhout,2 Terry J. Beveridge,1,2‡

John R. Dutcher,1,3 and Joseph S. Lam1,2*Biophysics Interdepartmental Group,1 Department of Molecular and Cellular Biology,2 and

Department of Physics,3 University of Guelph, Guelph, Ontario, Canada N1G 2W1

Received 28 May 2009/Accepted 24 August 2009

Bacterial biofilms are responsible for the majority of all microbial infections and have profound impact onindustrial and geochemical processes. While many studies documented phenotypic differentiation and generegulation of biofilms, the importance of their structural and mechanical properties is poorly understood. Herewe investigate how changes in lipopolysaccharide (LPS) core capping in Pseudomonas aeruginosa affect biofilmstructure through modification of adhesive, cohesive, and viscoelastic properties at an early stage of biofilmdevelopment. Microbead force spectroscopy and atomic force microscopy were used to characterize P. aerugi-nosa biofilm interactions with either glass substrata or bacterial lawns. Using isogenic migA, wapR, and rmlCmutants with defined LPS characteristics, we observed significant changes in cell mechanical properties amongthese strains compared to wild-type strain PAO1. Specifically, truncation of core oligosaccharides enhancedboth adhesive and cohesive forces by up to 10-fold, whereas changes in instantaneous elasticity were correlatedwith the presence of O antigen. Using confocal laser scanning microscopy to quantify biofilm structuralchanges with respect to differences in LPS core capping, we observed that textural parameters varied withadhesion or the inverse of cohesion, while areal and volumetric parameters were linked to adhesion, cohesion,or the balance between them. In conclusion, this report demonstrated for the first time that changes in LPSexpression resulted in quantifiable cellular mechanical changes that were correlated with structural changesin bacterial biofilms. Thus, the interplay between architectural and functional properties may be an importantcontributor to bacterial community survival.

Biofilms are sessile microbial communities growing on asurface or at an interface, often enmeshed in polymeric sub-stances. Being the predominant mode of microbial growth innature, bacterial biofilms are particularly problematic in thecontext of human health, accounting for up to 80% of allbacterial infections. In industrial processes, bacterial biofilmscause corrosion and biofouling, resulting in considerable lossof productivity. In the natural environment, biofilms play a rolein modulating worldwide geochemical cycles. Given the impactof biofilms in these diverse areas, the need for developingeffective strategies to control them is of paramount impor-tance. Since bacterial cell surface structures are convenienttargets for control agents, their roles in influencing biofilmfunction and architecture warrant in-depth investigations. Todate, most studies of biofilms have focused on genetic regula-tion, phenotypic differentiation and their contribution to anti-biotic resistance. In contrast, the mechanical and structuralproperties that link the genotypes to phenotypes of bacterialbiofilms are not well understood and rarely studied in a quan-titative and correlated manner.

Pseudomonas aeruginosa is a gram-negative opportunisticpathogen implicated in serious infections in patients with cysticfibrosis and immunocompromised patients. This bacterium hasa relatively large genome (6.3 Mb) consistent with its propen-sity to utilize versatile metabolic pathways, thereby developingantibiotic resistance and producing an arsenal of virulencefactors, including lipopolysaccharide (LPS) present on the cellsurface. LPS is localized in the outer leaflet of the outer mem-brane of all gram-negative bacteria, forming the first point ofcontact between the bacterial cell and any surface that it col-onizes or therapeutic agents. The LPS of P. aeruginosa consistsof three regions: lipid A, core oligosaccharide (core OS), andO antigen. The O antigen is synthesized as two distinct formswith overlapping pathways: the shorter A-band homopolymeris the so-called “common polysaccharide antigen” among thisspecies and consists of repeating D-rhamnose (D-Rha) sub-units, while the longer B-band heteropolymer is composed ofrepeating tri- to pentasaccharide subunits that vary among the20 serotypes of P. aeruginosa (42). The core OS is conceptuallydivided into the highly conserved inner core and the morevariable outer core. Depending on the linkage of L-rhamnose(L-Rha) with two distinct D-glucose (D-Glc) residues, two mainglycoforms of the core OS exist (see Fig. 1A). In the “capped”glycoform, L-Rha is �-1,3 linked to a D-Glc and acts as theacceptor molecule for O antigen, resulting in the production ofsmooth LPS. In the “uncapped” glycoform, L-Rha is �-1,6linked to a different D-Glc and is not substituted with O anti-gen, resulting in the production of rough LPS (39). In addition,

* Corresponding author. Mailing address: Department of Molecularand Cellular Biology, University of Guelph, Guelph, Ontario, CanadaN1G 2W1. Phone: (519) 824-4120, ext. 53823. Fax: (519) 837-1802.E-mail: [email protected].

† Supplemental material for this article may be found at http://jb.asm.org/.

‡ T.J.B. is deceased.� Published ahead of print on 28 August 2009.

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the presence or absence of the �-1,6 linked L-Rha substitutedwith a terminal D-Glc gives rise to the so-called intact or trun-cated outer core, respectively. The functional significance ofthis terminal glucose is unclear at present, although a role inhost cell binding has been proposed (57).

Mechanical processes that are important in the biofilm lifecycle include bacterial adhesion, cohesion, and viscoelasticity.Bacterial adhesion is a prerequisite for surface colonizationand the most important functional determinant in the earlystages of biofilm development. Accurate measurement of ad-hesion is therefore essential for monitoring the tendency ofbacteria to attach to surfaces and to switch from a planktoniclifestyle to a biofilm lifestyle. Data accumulated in previousstudies suggest that LPS is involved in bacterial cell adhesionto both abiotic (2, 8, 20, 32, 35, 54, 56) and biotic (17, 38, 49,56, 57) surfaces. Moreover, environmental factors, such asgrowth temperature, pH, ionic strength, nutrient availability,and oxygen levels, may influence cell adhesion via modificationof LPS expression and conformation (16, 36, 46, 47, 53). Theeffect of LPS on bacterial adhesion to various types of surfacesapparently involves distinct and complex mechanisms that re-main to be elucidated.

Bacterial cohesion, herein defined as cell-to-cell adherence,is crucial to the formation of microbial flocs and the growthand detachment of established bacterial biofilms. Quantifica-tion of cohesion is important for understanding biofilm biol-ogy, and such data are crucial for modeling and forecastingbiofilm development so that better control strategies can bedeveloped (55). Previous studies of biofilm cohesiveness havecharacterized it as highly stratified (3, 4, 18, 43), influenced byionic strength (14, 34), proportional to shear rate (37), andoften variable over 3 orders of magnitude (40, 50). Althoughan earlier study by Spiers and Rainey (48) provided semiquan-titative measurements of the role of LPS on bacterial cohesionwithin a biofilm, a truly quantitative account of the effect ofLPS on biofilm cohesion has not been demonstrated.

Bacterial viscoelasticity refers to the combined liquid-likeand solid-like characteristics in the behavior of polymeric sys-tems such that when deformed under stress, their strain canincrease over time (i.e., creep) and their original shape may beonly partially restored upon stress relief (19). Although earlierreports suggested that LPS modulates bacterial cell compress-ibility and helps prevent catastrophic structural failure due tomechanical stress (1, 52), no direct physical evidence of itsinvolvement in these processes has yet been presented. There-fore, monitoring biofilm viscoelasticity is crucial for demon-strating how well biofilms resist stresses, due to, for instance,fluid shear and antimicrobial peptides (5, 6). To date, quanti-tative data on how LPS affects viscoelastic properties of bio-

films are lacking, and existing studies have merely focused onelasticity measurements (7, 52). Recently, our group has de-veloped an atomic force microscopy (AFM)-based techniquecalled microbead force spectroscopy (MBFS) to measure theadhesive forces and viscoelastic properties of cells within bac-terial biofilms (28). In this study, we expand the application ofthis MBFS method to measure cohesive forces between cells atan early stage of biofilm development.

Biofilm structure refers to the distribution of biomass orcarbonaceous materials associated with cells (including all vi-able and nonviable cells and their extracellular polymeric sub-stances) within the space occupied by a biofilm. It is known tobe very heterogeneous and highly stratified, typically composedof a cohesive basal layer and a relatively fragile top layer (15,18, 43). Using confocal laser scanning microscopy (CLSM),biofilm structure can be quantitatively described in terms oftextural and volumetric parameters (11, 29). Textural param-eters characterize the pattern of cell clusters and interstitialvoids in a biofilm, whereas volumetric parameters describe themorphological characteristics of bacterial biofilms in three di-mensions (3-D) (11). Biomass distribution is affected by thesurrounding environment and may reflect fundamental pro-cesses occurring within biofilms, such as nutrient transport,accumulation rate, microbial physiology, and mechanical be-havior (29). Therefore, quantifying biofilm structure by CLSMwill allow us to understand the underlying processes and therelationship between biofilm architecture and behavior (15).

To examine the effects of differential LPS core capping onthe mechanical properties of early biofilms (here defined asconfluent bacterial lawns that have just begun to develop intofull-fledged biofilms) and the structural properties of maturebiofilms, we compare P. aeruginosa wild-type strain PAO1 withthose of its isogenic migA, wapR, and rmlC mutant strains withdefects in the respective genes affecting LPS core biosynthesis(22, 31, 39, 41). The migA gene (PA0705) encodes the putative�-1,6-rhamnosyltransferase necessary for the attachment ofthe terminal D-Glc to the outer core (39). The wapR gene(PA5000) encodes the putative �-1,3-rhamnosyltransferasecrucial to the capping of the core with O antigen (39). ThermlC gene (PA5164) encodes a dTDP-4-dehydrorhamnose 3,5-epimerase essential in the biosynthesis of TDP–L-Rha, which isthe precursor for the L-Rha in the LPS core (41). Defects inmigA, wapR, and rmlC mutants result in the expression ofdifferent LPS phenotypes, including a truncated outer coreand/or a lack of capping by O antigen (Table 1). In this study,we test the hypothesis that LPS contributes to biofilm functionand architecture through modulation of cellular mechanics andmicrocolony structures, thereby contributing to bacterial com-munity survival. By correlating quantitative mechanical changes

TABLE 1. Pseudomonas aeruginosa strains used in this study

Strain Characteristic(s) LPS phenotypeb Reference

PAO1 P. aeruginosa wild-type strain; IATSa serotype O5 A�B�; capped, intact outer core Hancock and Carey (22)migA mutant PAO1-derived migA::Gmr knockout mutant A�B�; capped, truncated outer core Poon et al. (39)wapR mutant PAO1-derived wapR::Gmr knockout mutant A�B�; uncapped, intact outer core Poon et al. (39)rmlC mutant PAO1-derived rmlC::Gmr knockout mutant A�B�; uncapped, truncated outer core Lindhout et al. (31)

a IATS, International Antigenic Typing Scheme.b A�B�, devoid of A-band and B-band LPS; A�B�, presence of A-band and B-band LPS.

VOL. 191, 2009 LPS CORE AFFECTS BIOFILM STRUCTURE AND MECHANICS 6619

in early P. aeruginosa biofilms and structural changes in maturebiofilms due to differences in LPS chemistry, we aim to eluci-date how the properties of these important bacterial cell sur-face molecules can alter the physical nature of biofilms.

MATERIALS AND METHODS

Bacterial strains, growth, and harvesting. Pseudomonas aeruginosa wild-typestrain PAO1 and three isogenic mutant strains with differential LPS core cappingwere used in this study (Table 1). Bacteria were grown overnight (16 h) inTrypticase soy broth (TSB) at 37°C on an orbital shaker (125 rpm). The cellswere harvested by centrifugation at 2,300 � g for 5 min, and the pellets werewashed in sterile water or phosphate-buffered saline and recentrifuged. After thefinal resuspension, 10-fold dilutions were made, and the optical density at 600 nm(OD600) was measured. The original washed but undiluted cultures were ad-justed to the appropriate concentrations (see the individual descriptions of ex-periments below) for various experiments.

Biofilm growth in flow cells. Biofilms of P. aeruginosa strain PAO1 and itsthree isogenic mutants were grown in continuous-culture flow cells constructedof polycarbonate (Biosurface Technologies Inc., MT) with chamber dimensionsof 40 mm by 10 mm by 0.1 mm. Briefly, 75% (vol/vol) ethanol was first pumpedfrom a reservoir through silicone tubing at 1 ml/min using a peristaltic pump(Minipuls 2; Gilson, Inc., Middleton, WI) to sterilize the flow cells for at least12 h. Subsequently, 1/10 strength TSB (BD) was pumped from a 6-liter flask(medium reservoir) to as many as eight flow cells set up in parallel at a rate of1 ml/min for 2 h, before inoculation of washed and diluted overnight cultures(OD600 of 0.1) via upstream injection ports (while the line from the mediumreservoir was clamped).The flow of culture medium was suspended during inoc-ulation to facilitate adhesion and resumed 1 h postinoculation. The spent me-dium exiting the flow cells was collected in a 20-liter carboy (waste reservoir).Flow cells were operated for 3 days in experiments cultivating biofilms that wereused to isolate and characterize LPS and for 7 days in experiments cultivatingbiofilms that were evaluated by confocal microscopy (see below).

Preparation and analysis of LPS. Lipopolysaccharide was prepared by themethod of Hitchcock and Brown (23) and resolved by sodium dodecyl sulfate(SDS)-polyacrylamide gel electrophoresis. Briefly, overnight bacterial cultures orbacterial cells collected from disrupted 3-day-old biofilms were washed anddiluted to an OD600 of 0.5 in a volume of 1 ml per sample. Bacterial suspensionswere centrifuged at 2,300 � g for 5 min and resuspended in 250 �l of Hitchcockand Brown lysis buffer (2% SDS, 4% �-mercaptoethanol, 10% glycerol, 1 M TrisHCl [pH 6.8], 0.002% bromophenol blue). The samples were heated at 100°C for30 min, cooled to room temperature, and incubated with 1.5 �l of proteinase K(20-mg/ml stock; Roche Diagnostics, Mannheim, Germany) at 56°C for 2 h. Foreach sample, 5 �l of the preparation were resolved by electrophoresis on a 12.5%polyacrylamide gel at 150 V. After electrophoresis, polyacrylamide gels weresilver stained according to the ultrafast method of Fomsgaard et al. (21).

AFM imaging of bacteria. Topographic atomic force microscopy images wereobtained using an MFP-3D atomic force microscope (Asylum Research, SantaBarbara, CA). For imaging bacterial lawns, rectangular silicon cantilevers CSC38(no Al, type B) (Mikromasch USA, Wilsonville, OR) with manufacturer’s quotedresonance frequencies of �10 kHz (range, 7 to 14 kHz) and force constants of�0.03 N/m (range, 0.01 to 0.08 N/m) were used. Twenty microliters of washedand diluted overnight bacterial cultures (OD600 of 1.0) was deposited ontofreshly cleaved mica and air dried for 20 min. Atomic force micrographs werecollected by raster scanning the sample under contact mode in air with the setpoint adjusted to 0.2 V and the integral gain at 30 or under contact mode in waterwith the set point adjusted to 0.05 V and the integral gain at 50. Height,deflection, z-sensor and lateral signals were simultaneously collected for real-time image construction. The atomic force microscope was controlled using theMFP-3D software version 070111�217 (Asylum Research, Santa Barbara, CA)operating within the Igor Pro 6.02A software environment (Wavemetrics, Inc.,Lake Oswego, OR). The root mean square (RMS) roughness of cell surfaces wascalculated from height images after the background areas had been masked.Three-dimensional renditions of height images were constructed offline using theArgyle function in the MFP-3D program.

Microbead force spectroscopy. The MFP-3D atomic force microscope was alsoused in force mode to quantify the force of bacterial adhesion based on a methodcalled microbead force spectroscopy that was recently developed by our group(28). For quantitative measurement of forces, rectangular tipless silicon cantile-vers (CSC12 [tipless, no Al, type E]; Mikromasch USA, Wilsonville, OR) withthe manufacturer’s quoted resonance frequencies of �10 kHz (range, 7 to 14kHz) and force constants of �0.03 N/m (range, 0.01 to 0.08 N/m) were calibrated

in air by the thermal method (24) to derive an accurate spring constant for eachindividual cantilever. This was a two-step process involving measuring the inverseof the optical lever sensitivity by pressing on a hard surface and collecting thethermal nose power spectrum. Only cantilevers with a calibrated spring constantwithin the range of 0.015 to 0.045 N/m were accepted for use in force spectros-copy studies. Microsized glass beads with diameters of 50 �m (Polysciences, Inc.,Warrington, PA) were attached to the distal ends of the cantilevers with two-component epoxy glue using a micromanipulator and dried overnight. Beadedcantilevers were then coated with 0.01% poly-L-lysine by exposure for 1 min andair dried for 10 min. Overnight bacterial cultures that were washed and adjustedto an OD600 of 2.0 were applied to the poly-L-lysine-coated beads three times toensure confluence. After the cantilever was mounted onto the head of the atomicforce microscope and centering the infrared laser spot behind the bead, thecantilever assembly was immersed in 100 �l of sterile deionized water on aprecleaned glass slide or a mica surface precoated with a bacterial lawn as wasdone for imaging studies. The microbead was lowered gradually to approach theglass surface or the cell layer, upon which force curves (force-separation plots)and creep curves (indentation-time plots) were gathered simultaneously. In eachindividual experiment, 10 force curves and 10 creep curves were collected foreach sample under standard conditions, defined as a loading force of 10 nN,contact time of 1 s, ramp velocity of 2 �m/s, and ramp distance of 3 �m (28).

Fluorescence labeling and confocal laser scanning microscopy. After 7 days ofgrowth, biofilms in flow cells were stained in situ using reagents from the Live/Dead BacLight bacterial viability kit (BD Biosciences). Equal proportions of thegreen SYTO 9 stain for live cells and the red propidium iodide stain for deadcells were mixed and diluted in sterile deionized water (3:1,000), and 1 ml of thismixture was injected into the upstream port of each flow cell. Biofilms wereincubated in complete darkness for 30 min (which was found to be sufficient timefor stain penetration), and unbound stains were removed by injection of 3 ml of1/10 strength TSB into each upstream port. Imaging by CLSM was performedimmediately and completed within 24 h using a confocal microscope (model TCSSP2; Leica Microsystems Canada Inc., Richmond Hill, Ontario, Canada). Foreach of the four bacterial strains investigated, optical sectioning was performedfor three representative microcolonies to obtain image stacks for structuralquantitation. Excitation wavelengths were set at 488 nm and 543 nm, whileemission bandwidths were set at 500 to 535 nm and 555 to 700 nm, respectively,for detecting the green and red channels. Image stacks consisted of image slicesspaced 1 �m apart, starting from the substratum and ending at the top ofmicrocolonies. Image slices were gathered at a scan speed of 400 Hz and aresolution of 1,024 by 1,024 pixels using a Leica PL Fluotar 40.0 � 1.00 oilimmersion objective lens (with the 10� eyepiece and 2� digital zoom, a totalmagnification of �800). The microscope was controlled using the proprietaryLeica confocal software, a platform in which data for cross sections and averageprojections of microcolonies could be obtained.

Quantitation of biofilm structure. Biofilm structural quantitation was per-formed on the CLSM image stacks by the methods of Lewandowski and Beyenal(29). In such 3-D analyses, image stacks were exported as individual images inTIFF format from the Leica application suite advanced fluorescence lite pro-gram (Leica) into XnView (freeware by Pierre Gougelet, available at http://www.xnview.com), batch converted to gray-scale images, and resized to 188 by 188pixels such that “voxel” dimension was 1 �m by 1 �m by 1 �m. Using the ISA-3Dpackage for biofilm image analysis (Center for Biofilm Engineering, MontanaState University, Bozeman, MT) within the MATLAB and Image Analysis Tool-box environments (Mathworks, Inc., Natick, MA), textural parameters (i.e.,textural entropy, energy, and homogeneity) were extracted from the gray-scaleimages, whereas volumetric parameters (i.e., average run lengths, aspect ratios,diffusion distances, fractal dimension, porosity, biovolume, biomass and biofilmthicknesses, biofilm and biomass roughnesses, biomass surface area, and biovol-ume-to-biomass surface area ratio) were calculated from binary images after anautomatic threshold protocol was applied.

Contact angle measurements. For the measurement of contact angles, a sessiledrop technique based on the method of Korenevsky and Beveridge was per-formed (27). Briefly, overnight cultures of P. aeruginosa cells were harvested andwashed twice with a sterile buffer (0.1 M NaCl–0.05 M HEPES [pH 7.4]) andadjusted to an OD600 of 1.0 in a 10-ml volume of the buffer. The bacterial cellswere then deposited on cellulose acetate membrane filters (AcetatePlus, sup-ported, plain, 0.22 micron, 47 mm; GE Water & Process Technologies) viavacuum aspiration to produce an even, confluent bacterial lawn. Bacterial layerswere dried for 30 min before the so-called “plateau contact angles” were mea-sured using sterile deionized water droplets. Water droplets of approximately 2�l were deposited onto the bacterial lawns using a micropipette (Gilson, Inc.,Middleton, WI). To obtain static contact angles, water droplets were allowed tosettle for 2 s before digital images were captured from the side using a 12.1-

6620 LAU ET AL. J. BACTERIOL.

megapixel charge-coupled-device camera (Sony of Canada Ltd., Toronto, On-tario, Canada) operating in macro mode, against a bright illuminating back-ground. Saved images were processed by a program (designed by Joop der Vries,University of Groningen, The Netherlands) based on the approximation thatsessile droplets form spherical caps. Mean contact angles for each sample werecalculated from five different droplets deposited on different areas of the mem-brane, with five replicate measurements per droplet.

BATH assay. The relative hydrophobicity of cells was examined using the bacte-rial adhesion to hydrocarbon (BATH) assay (45). Briefly, cells cultured in LB brothwere standardized to an OD600 of 0.3, harvested by centrifugation at 5,000 � g, and

washed once in 0.1 M potassium phosphate buffer (pH 7.0) before resuspension in1.5 ml of the same buffer. The cell suspension (1.2 ml) was transferred to a glass testtube (10-mm diameter) and incubated with 200 �l of hexadecane (Fisher Scientific)for 10 min at 30°C. The cell and hexadecane mixture was vigorously mixed by usinga Vortex (VWR Scientific) for a total time of 2 min, followed by further incubationfor 25 min at room temperature. The OD595 of the aqueous layer (ODaq) wasmeasured on a BMG FluoStar Optima plate reader spectrophotometer and com-pared to the OD595 of the cell suspension before the addition of hexadecane (ODcs)in order to determine the percent adherence using the following equation: percentadherence � [1 � (ODaq/ODcs)] � 100.

FIG. 1. Comparison of LPSs from Pseudomonas aeruginosa wild-type strain PAO1 and mutant strains with LPS core variants. (A) Schematicdiagram illustrating the chemical structures of smooth LPS and rough LPS. The gray and black arrows point to the position of outer core truncationand position of O-antigen capping, respectively. (B) Silver-stained SDS-polyacrylamide gels illustrating LPS profiles of planktonic and biofilm cells.The two gray arrows and the black arrows point to the position of O antigen and position of core-plus-one entities, respectively. Planktonic cells(lanes 1 to 4) and biofilm cells (lanes 5 to 8) of strain PAO1 (P), migA mutant (M), wapR mutant (W), and rmlC mutant (R) are shown.


Bacterial clumping assay. The clumping and resultant precipitation of bacte-rial cells were monitored and quantified over a 24-h period as follows. Eachstrain of P. aeruginosa was grown overnight (16 h) in 10 ml of TSB in glass testtubes at 37°C with shaking. The cultures were then placed upright in a test tuberack to allow flocculation to proceed. Finally, at 0, 1, 2, 3, 4, and 24 h after thebeginning of the assay, 100-�l aliquots were taken from near the surface (5-mmdepth) of each culture and diluted 10-fold for OD600 measurements.

RESULTS

The P. aeruginosa wild-type strain PAO1 and migA mutantstrain showed differences between planktonic and biofilm LPS.The three isogenic mutants of P. aeruginosa PAO1 investigated inthis study, namely, the migA, wapR, and rmlC mutants, havedifferential core capping in their LPS moieties (Fig. 1A and Table1). Silver-stained SDS-polyacrylamide gels of LPS from plank-tonic cells revealed that the rough strains lacking LPS O antigen(i.e., wapR and rmlC mutants) are devoid of long-chain O poly-

saccharides as well as core OS capped with one O-antigen repeatunit, i.e., the “core-plus-one” entity (Fig. 1B). Interestingly, thebiofilm cells of these strains appeared to have LPS profiles similarto those of their planktonic counterparts. In contrast, the PAO1strain and migA mutant, which both possess O polysaccharides,showed different profiles between planktonic and biofilm LPS.Specifically, the LPS bands at intermediate-molecular-weight Oantigen present in the LPS sample from planktonically grownPAO1 cells were absent in LPS prepared from the same straingrown in biofilm mode. These intermediate-molecular-weightbands were also absent in LPS samples prepared from the migAmutant grown planktonically as well as in migA cells grown asbiofilm cultures in the flow chamber. Furthermore, production ofboth the full-length O antigen and the core-plus-one entitypresent in the former sample were significantly reduced in thelatter sample, likely due to downregulation.

FIG. 2. Atomic force micrographs of Pseudomonas aeruginosa wild-type strain PAO1 and mutant strains with LPS core truncations collectedin contact mode in air. (A) Strain PAO1; (B) migA mutant; (C) wapR mutant; (D) rmlC mutant. Height images (top panels) are shown withenclosed areas for roughness calculations. Deflection images (bottom panels) reveal more details in cell morphology. The straight lines in panelA are steps in the mica substratum.


Atomic force micrographs revealed differential surface mor-phology of P. aeruginosa wild-type strain PAO1 and mutantstrains with LPS core variants. Atomic force micrographs ofP. aeruginosa cells scanned in air revealed changes in cellsurface morphology in the LPS truncation mutants comparedto the wild type (Fig. 2). While deflection images were usefulfor the emphasis of textural details, height images were ob-tained for the calculation of surface roughness after masking toeliminate background areas (Fig. 2A to D). The values forRMS roughness for strain PAO1 (wild type) and migA, wapR,and rmlC mutants were found to be 49.43 nm, 68.41 nm, 44.40nm, and 34.14 nm, respectively. Images collected in aqueousmedium did not show any discernible differences in appear-ance among the cells of the different strains (data not shown).Three-dimensional rendering of the height images in air wereproduced to highlight changes in surface topography (data notshown). The textures of cells shown in these images corre-sponded with RMS roughness data and further demonstratedthat bacterial cells from strains possessing O antigen have

rougher cell surfaces, while strains devoid of O antigen have asmoother appearance.

MBFS data quantitatively demonstrated enhanced cell ad-hesion, cohesion, and altered viscoelasticity in early biofilmsof mutants with LPS core variants. Using microbead forcespectroscopy with two different bead and surface configura-tions (see Fig. S1 in the supplemental material), we were ableto precisely quantify the forces of early biofilm adhesion toglass and cell-cell cohesion (see Fig. S2A in the supplementalmaterial) for the wild-type strain PAO1 and its three isogenicmutants. Overall, cell adhesion and cohesion were enhanced inthe LPS core truncation mutants than in the wild type (Fig. 3).The average adhesive force of bacteria to glass measured forwild-type strain PAO1 and migA, wapR, and rmlC mutantsunder standard conditions were 0.66 0.27 nN, 5.52 1.83nN, 6.90 1.22 nN, and 5.12 1.12 nN, respectively (Fig. 3A).The corresponding average cell-cell cohesive forces for strainPAO1 and migA, wapR, and rmlC mutants were 1.00 0.31nN, 2.99 0.13 nN, 5.15 1.09 nN, and 12.18 0.15 nN,

FIG. 2—Continued.


respectively (Fig. 3B). The inverses of adhesion and cohesionas well as their ratios were plotted to reveal other possibletrends when comparing the four bacterial strains to each other(see Fig. S3 in the supplemental material). These trends al-lowed the assessment of correlation between quantitativechanges in mature biofilm structure with variations in earlybiofilm mechanics.

For the determination of early biofilm viscoelastic proper-ties, fitting of data from the MBFS experiments to a Voigt

standard linear solid model (28) yielded values for instanta-neous and delayed elastic moduli, viscosity, and characteristicresponse time (i.e., retardation time) (see Fig. S2B in thesupplemental material). The viscoelastic properties of allthe LPS core truncation mutants were different from thoseof the wild type (Fig. 4). Instantaneous elastic moduli werefound to be larger for wild-type strain PAO1 and migAmutant (1.69 � 105 Pa 0.80 � 105 Pa and 1.88 � 105 1.08 � 105 Pa, respectively) than for wapR and rmlC mutantstrains (6.25 � 104 1.90 � 104 Pa and 7.96 � 104 2.38 �104 Pa) (Fig. 4A). Delayed elastic moduli were larger for thePAO1 strain (1.09 � 106 0.10 � 106 Pa) than for the migA,wapR, and rmlC mutants (4.78 � 105 0.61 � 105 Pa,6.12 � 105 4.60 � 105 Pa, and 7.48 � 105 6.15 � 105 Pa,respectively) (Fig. 4B). The viscosity values for strain PAO1and migA, wapR, and rmlC mutants were found to be 2.31 �105 1.06 � 105 Pa � s, 1.28 � 105 0.05 � 105 Pa � s,2.12 � 105 1.66 � 105 Pa � s, and 3.72 � 105 3.25 � 105

FIG. 3. Biofilm adhesive and cohesive forces of Pseudomonasaeruginosa wild-type strain PAO1 and mutants with LPS core variantsas measured by MBFS. Wild-type strain PAO1 and migA, wapR, andrmlC mutant strains were examined. Averages of cell-glass adhesion(A) and cell-cell cohesion (B) measurements from three independentexperiments are shown. Error bars denote standard errors of measure-ment.

FIG. 4. Bacterial viscoelastic properties for Pseudomonas aeruginosa wild-type strain PAO1 and mutants with LPS core variants as measuredby MBFS. Wild-type strain PAO1 and migA, wapR, and rmlC mutant strains were examined. (A) Instantaneous elastic modulus; (B) delayed elasticmodulus; (C) viscosity; (D) characteristic response time. Error bars indicate standard errors of measurement.

FIG. 5. Differential clumping and precipitation of Pseudomonasaeruginosa wild-type PAO1 and mutants with LPS core variants. Wild-type strain PAO1 and migA, wapR, and rmlC mutant strains wereexamined. Results are shown from three independent experiments.


Pa � s, respectively (Fig. 4C), whereas the characteristic re-sponse times were measured and found to be 0.204 0.078s, 0.274 0.046 s, 0.327 0.026 s and 0.432 0.079 s,respectively, for these strains (Fig. 4D).

Bacterial clumping observed corresponded with cell-cell co-hesion measurements by MBFS. Bacterial cultures of LPS coremutants exhibited a higher degree of clumping than those ofthe wild-type strain (see Fig. S4 in the supplemental material).Previous work carried out by our group also indicated that thelarger the truncation of the core OS and/or the O antigen, thegreater the resultant level of cell clumping in bacterial biofilmsgrown on glass slides (30). We therefore carried out semiquan-titative bacterial clumping assays to verify these observations.Cells in broth cultures of strain PAO1 and migA, wapR, andrmlC mutants exhibited increasing tendencies, in that order, toflocculate (form clumps) and settle at the bottom of stationarytest tubes (Fig. 5). These observations showed good correlationwith quantitative measurements of cell-cell cohesion forces forthese respective bacterial strains (Fig. 3B).

BATH assays corresponded with contact angle measure-ments. The bacterial adhesion to hydrocarbon assay, thoughindirect, has been routinely used for examining the hydropho-bicity of cells and is based on the fact that cells with morehydrophobic surfaces will adhere more readily to a droplet ofhydrocarbon (44). Under the standardized experimental con-ditions described in Materials and Methods, cells of the wild-type strain PAO1 and the migA, wapR, and rmlC mutants withLPS variants adhered to n-hexadecane with increasing affinity,with 33.5%, 38.2%, 40.2%, and 45.2% of cells adhered onaverage, respectively (Table 2). These data should theoreticallycorrespond with contact angle measurements between waterand bacterial lawns using a sessile drop technique, since con-tact angles are characteristic of interfacial energies and thusalso dependent on cell surface hydrophobicity (51). Accord-ingly, bacterial lawns of the wild-type strain PAO1 and itsisogenic LPS migA, wapR, and rmlC mutants were shown tocontact water droplets in an increasing trend with angles of33.3°, 34.0°, 34.1°, and 37.5°, respectively (Table 2).

Differential LPS core capping results in quantifiablechanges in mature biofilm structure that could be correlatedto early biofilm mechanics. Confocal micrographs of the bac-terial strains examined showed that while cells of wild-typestrain PAO1 form microcolonies with round perimeters, themicrocolonies formed by the LPS mutants have more irregularedges (Fig. 6). Interestingly, biomass thickness was found to beconsistently higher for strain PAO1 and rmlC mutant. Thesequalitative observations were confirmed by quantitative com-parison of average run lengths and biomass thicknesses, re-spectively (Table 3). Quantitative biofilm structure analysis of

microcolony image stacks in 3-D also indicated that there weresignificant changes of textural and volumetric parameters be-tween wild-type strain PAO1 and its isogenic mutants thatcould be correlated with differences in force and viscoelasticitymeasurements (Table 4) (see Fig. S5 in the supplemental ma-terial).

DISCUSSION

Until this study, the contribution of LPS to the mechanicalproperties of bacterial biofilms had not been defined. The fewexisting quantitative studies of how LPS affects bacterial adhe-sion have given inconclusive results. For example, Burks et al.(13) performed nanoscale adhesion measurements by conven-tional force spectroscopy on three E. coli strains of differingLPS phenotypes but were unable to correlate the results withdata from macroscale assays they gathered based on binding toglass bead columns. Likewise, Abu-Lail and coworkers (2)observed that bacterial adhesion to silica is significantly higherfor P. aeruginosa strain AK1401, a mutant that lacks B-band Oantigen, than for P. aeruginosa PAO1, while adhesion to or-ganics is stronger in the wild-type strain. In separate studies,however, this research group observed that adhesion to siliconwas stronger for strain PAO1 than for strain AK1401 (8), whileadhesion to serum albumin was three times higher in the mu-tant (9). These conflicting results in comparing the adhesivestrengths of wild-type and LPS variant strains might have beencaused by the use of sample preparation techniques and testingenvironments that have not been standardized in the differentstudies (28). To address these issues, we employed the MBFSmethod to measure the mechanical properties of P. aeruginosacell layers derived from strains with different LPS phenotypesand further correlated the differences in mechanical behaviorat this early stage of biofilm formation to matrix architecturalchanges in fully developed biofilms, as quantified by imageanalysis of confocal laser scanning micrographs.

Although bacterial cells growing in a biofilm have beenshown to lose B-band O antigen from their LPS (10), the roleof core OS structure in this process is unclear. The LPS-banding profiles of samples prepared from strain PAO1 and itscore-deficient mutants in SDS-polyacrylamide gels clearly re-vealed that strain PAO1 and migA mutant had substantiallydifferent patterns when grown planktonically and in a biofilm,albeit in different ways. In strain PAO1, the medium-length Oantigen present in LPS prepared from planktonically growncells was absent when the wild-type strain was grown as bio-films, whereas in the migA mutant, no medium-length O anti-gen could be discerned in the LPS prepared from planktoni-cally grown cells. In addition, O antigen and “core-plus-one”bands were also absent from LPS samples of migA mutantgrown as biofilms. These observations suggested that the outercore defect caused by mutation in the migA mutant might leadto instability or downregulation of O-antigen capping. A moreintriguing possibility is that core OS integrity may have beencompromised, as suggested by the appearance of a new bandrunning slightly faster than core OS in the migA mutant biofilmsample (Fig. 1B).

Bacterial cells with and without LPS O antigen generallyexhibit “smooth” and “rough” colony morphology of bacterialgrowth, respectively. Interestingly, high-resolution images of

TABLE 2. Comparison of bacterial adhesion to hydrocarbon assaydata with bacterial contact angle measurements obtained usingthe sessile drop technique on confluent lawns of P. aeruginosa

Strain % Adhesion in the BATHassay (mean SD)

Contact angle (°)(mean SD)

PAO1 33.5 9.1 33.3 2.8migA mutant 38.2 6.0 34.0 6.0wapR mutant 40.2 11.5 34.1 7.5rmlC mutant 45.2 9.8 37.5 3.6


strain PAO1 and its LPS core variants captured by AFMshowed that the roughness of the surface of individual bacterialcells is inversely related to colony roughness. The “smooth”strains possessing O antigen exhibited rough cell surfaces,likely due to the presence of higher proportion of exopolysac-charide, while “rough” strains without the O polysaccharideshowed smoother cell surfaces. Also, the observation that themigA mutant strain has rougher cell surfaces than strain PAO1(as measured by the RMS roughness) is logical because theloss of O antigen from some migA mutant LPS can lead to amix of smooth and rough LPS, resulting in greater variations insurface topography. Despite these differences, the contributionof cell surface roughness to bacterial interaction with externalsurfaces remains unclear, since surface roughness simulta-neously increases friction and reduces contact area, two factorsthat have opposing effects on adhesion. Finally, although dry-ing time had been standardized for AFM imaging, differentialcontribution of drying artifacts in the various strains to RMSroughness measurements could not be ruled out.

Bacterial adhesion is a fundamental requirement for cells to

attach to substrata for biofilms to form. Measurements of bac-terial adhesion at an early stage of biofilm formation usingMBFS showed that strains with rough LPS (wapR and rmlCmutants) have quantitatively stronger adhesion to glass thanstrains with smooth LPS do (strain PAO1 and migA mutant).These results are consistent with data from assays that mea-sured bacterial adhesion to glass that were previously per-formed by our group, suggesting that increasing defects in theLPS core and O antigen lead to an increasing degree of adhe-sion to glass (30). Intriguingly, the trend in adhesive forcemeasured by MBFS (strain PAO1 migA mutant wapRmutant � rmlC mutant) was not monotonically increasing, withthe wapR mutant having a stronger adhesion to glass than thermlC mutant strain did. This trend contrasted with that foundin our earlier study performed using planktonic cells and sug-gested a role for the terminal D-glucose in the intact core OSof the wapR mutant in strengthening adhesion to glass withinthe context of a biofilm.

Bacterial cohesion is a critical link between adhesion andviscoelasticity. Force measurements by MBFS showed that

FIG. 6. Confocal laser scanning micrographs illustrating the changes in biofilm structure resulting from differential LPS core capping. Averageprojections (top panels) and midpoint cross sections (bottom panels) of representative microcolonies of wild-type strain PAO1 (A), migA mutant(B), wapR mutant (C), and rmlC mutant (D) are shown.


cells with increasing core and O-antigen defects exhibit in-creasing cell-cell cohesion. As expected, this trend in cohesiveforce (strain PAO1 migA mutant wapR mutant rmlCmutant) correlated well with visual observations and semi-quantitative assays of bacterial clumping. In addition, cohesionmay also be linked to cell surface hydrophobicity as measuredby bacterial adhesion to hydrocarbons and by contact angles.However, the minor variations in hydrophobicity compared tothe much larger differences in cohesion among the four strainstested suggest that hydrophobicity played only a minor role incell-cell cohesion.

The balance between bacterial adhesion and cohesion iscrucial to understanding the structural development of a bio-film. For instance, biofilm adhesive and cohesive strengths maycontribute to the differential effects of solid surface tension,hydrodynamic shear, and disinfecting agents on different bio-films (12, 33). Remarkably, certain trends in adhesion andcohesion and their derived quantities (i.e., inverses and ratios)among the four tested P. aeruginosa strains proved to be highlycorrelated with some viscoelastic properties and quantitativestructural data (see below).

Bacterial viscoelasticity is an important determinant of howbacteria growing as biofilms respond to physical stress. Sincebiofilms are known to be viscoelastic materials that exhibitboth viscous and elastic behaviors, we modeled them using asimple mechanical analog (the Voigt standard linear solidmodel) consisting of a spring in series with a spring-dashpotelement in parallel (28). Measurements of biofilm viscoelastic-ity of the bacterial strains used in this study showed that fourmain trends exist: (i) link to the presence of O antigen (strainPAO1 � migA mutant � wapR mutant � rmlC mutant), asseen for instantaneous elastic modulus; (ii) correlation withinverse adhesion (PAO1 � migA mutant � wapR mutant rmlC mutant), as seen for delayed elastic modulus (correlationcoefficient [R] � 0.9119); (iii) correlation with the cohesion-to-adhesion ratio (PAO1 � migA mutant wapR mutant rmlC mutant), as seen for viscosity (R � 0.9496); and (iv)correlation with cohesion (PAO1 migA mutant wapRmutant rmlC mutant), as seen for characteristic responsetime (R � 0.9799) (see Fig. S6 in the supplemental material).Thus, the presence of O antigen appeared to be important inthe immediate response of a biofilm to mechanical stress

FIG. 6—Continued.


(instantaneous elasticity). In contrast, inverse adhesion in-fluenced the continued response to sustained indentation(delayed elasticity). Moreover, the ratio of cohesion overadhesion was important for determining the rate of irrevers-ible deformation (viscosity). Finally, cohesion affected theinitial interval in which rapid creep (�63% of total creep)occurs after a constant stress is applied, a characteristicresponse time derived by dividing viscosity over delayedelasticity (28).

Biofilm structure can be defined as the spatial distribution ofbiomass within a biofilm. The most practical approach forquantifying biomass distribution is by analyzing confocal mi-croscopy images and taking into consideration parameters thatcharacterize biofilm structures. Since these parameters areonly mathematical functions (with arbitrary units) characteriz-ing pixel distribution in the biofilm images, they must be cho-sen carefully for their relevance to the underlying biofilm pro-cesses. Following the method of Lewandowski and Beyenal

(11, 29), we calculated selective parameters that are deemeduseful for describing biofilm structure, and we further corre-lated these parameters to mechanical properties of cell layersat an early stage of biofilm formation (see Fig. S7 in thesupplemental material). When the confocal image stacks ofrepresentative microcolonies were subjected to structuralquantification, 3-D textural parameters and volumetric param-eters were derived. Trends for textural parameters correlatedeither with cell adhesion, as seen for energy (R � 0.9477) andhomogeneity (R � 0.9701), or with inverse cohesion, as seenfor textural entropy (R � 0.9327). The volumetric parametersfollowed five different trends: (i) correlation with adhesion,such as for average x and y run lengths (R � 0.7358 and 0.7227,respectively), average and maximum diffusion distances (R �0.8965 and 0.9715, respectively), average biofilm roughness(R � 0.8982), and biovolume-to-biomass surface area ratio(R � 0.7605); (ii) correlation with inverse adhesion, as forfractal dimension (R � 0.9945); (iii) correlation with cohesion,

TABLE 3. Quantitative changes in 3-D biofilm structure resulting from differential LPS core cappinga

Structural parameterbValue (mean SE) for:

PAO1 strain migA mutant wapR mutant rmlC mutant

TexturalTextural entropy 7.14 0.21 6.62 0.26 5.98 0.52 6.09 0.54Energy 0.01 0.00 0.02 0.01 0.04 0.02 0.03 0.01Homogeneity 0.32 0.02 0.43 0.03 0.51 0.05 0.46 0.04

VolumetricAvg run lengths (�m)

AXRL 7.54 0.61 18.05 4.24 12.77 0.96 15.90 8.21AYRL 7.40 0.60 17.56 4.57 12.18 0.86 14.53 7.54AZRL 7.78 0.78 12.53 1.80 10.23 1.40 14.49 6.01

Aspect ratiosAXRL/AYRL 1.02 0.01 1.04 0.03 1.05 0.01 1.09 0.03AXRL/AZRL 0.98 0.08 1.42 0.21 1.32 0.25 1.01 0.12AYRL/AZRL 0.96 0.07 1.37 0.22 1.25 0.22 0.94 0.11

Diffusion distances (�m)Avg 2.38 0.16 4.13 0.61 4.14 0.33 4.01 1.53Maximum 17.11 2.74 22.49 5.13 21.98 2.61 19.51 4.86

Fractal dimension 2.71 0.01 2.54 0.03 2.51 0.10 2.52 0.09

Porosity 0.65 0.03 0.62 0.05 0.70 0.09 0.66 0.09

Biovolume (� 105 �m3) 6.21 0.81 5.42 0.27 5.87 3.53 8.64 2.69

Thicknesses (�m)Biomass avg 46.60 3.79 38.68 2.37 44.68 13.97 63.28 6.78Biomass maximum 50.33 4.41 40.33 3.33 47.00 15.04 70.00 5.13Biofilm avg 0.10 0.02 0.04 0.01 0.07 0.02 0.14 0.04Biofilm maximum 40.47 2.99 29.72 4.78 34.13 19.09 51.45 8.56

RoughnessesBiomass 50.33 4.41 40.33 3.33 47.00 15.04 70.00 5.13Biofilm 0.31 0.03 0.49 0.14 0.71 0.32 0.46 0.13

Biomass surface area (� 105 �m2) 3.52 0.45 1.75 0.18 2.36 1.47 3.19 1.08

Biovolume-to-biomass surfacearea ratio (�m)

1.77 0.09 3.19 0.50 2.57 0.18 3.09 1.30

a Image stacks of three representative microcolonies were analyzed per strain.b AXRL, average x run length; AYRL, average y run length; AZRL, average z run length.


such as for average z run length (R � 0.8122) and aspect ratioxy (R � 0.9932); (iv) correlation with the adhesion-to-cohesionratio, such as for aspect ratios xz and yz (R � 0.9691 and0.9899, respectively); and (v) correlation with the cohesion-to-adhesion ratio, such as for biovolume (R � 0.9475), biomassand biofilm thicknesses (R � 0.9514 and 0.9937, respectively),average biomass roughness (R � 0.9908), and biomass surfacearea (R � 0.8020). On the basis of the above analyses, wewere able to conclude that in general, textural parameters weremost related to early biofilm adhesion, whereas volumetricparameters were variously linked to adhesive and cohesiveproperties and the balance between them. In a conceptualmodel linking mechanical and structural properties, it is rea-sonable to assume that a biofilm in a flowing environmentgrows thicker if cohesion is stronger than adhesion but be-comes more spread out over a surface if adhesion predomi-nates. This conjecture has indeed been proven correct byour observations that the wild-type strain PAO1 and rmlCmutant have both higher cohesion-to-adhesion ratios andthicker biofilms than the migA and wapR mutants do. Inter-estingly, the only structural parameter not correlated withforce measurements was porosity. Since this parameter,which had previously been related to biofilm accumulationrate (29), was instead linked to the presence of an intactouter core in this study, our observation implicates the in-tegrity of the LPS outer core in optimal metabolic activity.For a more detailed description of the biofilm structuralquantification parameters, interested readers are referred toTable S1 in the supplemental material.

While unicellular organisms can simply modulate theirstructure to bring about a functional response to an external

stimulus, the coordinated responses of multicellular systemssuch as bacterial biofilms to environmental stresses aremuch more complex. Specifically, structural changes in in-dividual cells need to be translated into mechanical andarchitectural modifications in the entire cell assembly forresponses to be effective. Previous studies linking biofilmmechanics and structures (25, 26) merely provided qualita-tive observations and conceptual assessments of the prop-erties of biofilms but did not take into account the roles ofspecific cell surface features such as LPS expression. In thisstudy, we measured the mechanical properties in the earlybiofilms of P. aeruginosa wild-type strain and mutants withLPS core variants and related them to quantitative struc-tural data in mature biofilms. The evidence presented heresuggests that the structure of bacterial biofilms is moststrongly correlated to the strength of adhesion, the balancebetween adhesion and cohesion, and the structural integrityof the LPS outer core OS. For the first time, changes inbacterial mechanical behavior during an early stage of bio-film formation caused by differential LPS core capping havebeen correlated with matrix architecture modifications infully developed biofilms. Also, the presence of a terminalglucose in the LPS core of P. aeruginosa appeared to belinked to core OS stability, increased adhesiveness in theabsence of O antigen, and optimal structural developmentwithin a biofilm. Evidently, LPS expression and the resultantinterplay between biofilm mechanics and structure may con-tribute importantly to bacterial community survival. There-fore, further investigation correlating these properties inother model microbial systems under various environmental

TABLE 4. Correlation of biofilm structural parameters with mechanical properties in Pseudomonas aeruginosa

Structural parameter Trenda Correlated property Correlationcoefficient

TexturalTextural entropy P � M � W R Inverse cohesion 0.9327Energy P M W � R Adhesion 0.9477Homogeneity P M W � R Adhesion 0.9701

VolumetricAvg x run length P M W � R Adhesion 0.7358Avg y run length P M W � R Adhesion 0.7227Avg z run length P M W R Cohesion 0.8122Aspect ratio xy P M W R Cohesion 0.9932Aspect ratio xz P M � W � R Adhesion-to-cohesion ratio 0.9691Aspect ratio yz P M � W � R Adhesion-to-cohesion ratio 0.9899Avg diffusion distance P M W � R Adhesion 0.9715Maximum diffusion distance P M � W � R Adhesion 0.8965Fractal dimension P � M � W R Inverse adhesion 0.9945Porosity P � M W � R �b �b

Biovolume P � M W R Cohesion-to-adhesion ratio 0.9475Avg biomass thickness P � M W R Cohesion-to-adhesion ratio 0.9514Maximum biomass thickness P � M W R Cohesion-to-adhesion ratio 0.9607Avg biofilm thickness P � M W R Cohesion-to-adhesion ratio 0.9937Maximum biofilm thickness P � M W R Cohesion-to-adhesion ratio 0.9607Biomass roughness P � M W R Cohesion-to-adhesion ratio 0.9908Biofilm roughness P M W � R Adhesion 0.8982Biomass surface area P � M W R Cohesion-to-adhesion ratio 0.8020Biovolume-to-biomass surface area ratio P M W � R Adhesion 0.7605

a P, PAO1 strain; M, migA mutant; W, wapR mutant; R, rmlC mutant. The trend shown in italic font does not conform to the trend of the most correlated property.b �, no strong correlation with mechanical properties could be found (correlation coefficient of 0.5).


stimuli will be of tremendous value to microbiological re-search.

ACKNOWLEDGMENTS

This work is supported by AFMNet funding and Discovery Grants toT.J.B. and J.R.D. from the Natural Sciences and Engineering Councilof Canada and an operating grant to J.S.L. from the Canadian CysticFibrosis Foundation. T.J.B. was a Canada Research Chair in the Struc-ture of Microorganisms, J.R.D. is a Canada Research Chair in SoftMatter Physics, and J.S.L. holds a Canada Research Chair in CysticFibrosis and Microbial Glycobiology.

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Differential Lipopolysaccharide Core Capping Leads to ...and architecture through modulation of cellular mechanics and microcolony structures, thereby contributing to bacterial com-munity