Impact of nano-topography on bacterial attachment

BiotechnologyJournal DOI 10.1002/biot.200700244 Biotechnol. J. 2008, 3, 536–544

536 © 2008 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim

1 Introduction

The mechanisms of bacterial attachment to sur-faces have been the focus of intense research overthe last few decades [1–10]. Theoretical approach-es such as the DLVO and thermodynamic theorieshave revealed some of the physico-chemical as-pects of bacterial adhesion, such as the influence of

surface charge and surface tension on the long-range cell-substratum interactions and the effect ofsurface hydrophobicity in the short-range interac-tions [11–15]. However, the bacterial attachmentprocess has a complex dependence on a number offactors relating to both the substratum and bacter-ial cell surface properties [3, 4, 7, 16].

For example, bacterial characteristics such ascell shape and size, production of extracellularpolymeric substances (EPS), surface protrusionssuch as flagella and cilia, cell-cell signaling, surfacetension and surface charge and wettability in par-ticular are believed to have a significant influenceon the process [17–23]. Although the wettabilityand charge characteristics of the cell surface bothresult from the presence and composition of differ-ent surface proteins, it is possible for the configu-ration of these surface molecules to change inphysiological response to environmental perturba-tions. This may result in different attachment be-

Research Article

Impact of nano-topography on bacterial attachment

Natasa Mitik-Dineva1, James Wang2, Radu C. Mocanasu1, Paul R. Stoddart2, Russell J. Crawford1 andElena P. Ivanova1

1Faculty of Life and Social Sciences, Swinburne University of Technology, Hawthorn, Australia2Center for Atom Optics and Ultrafast Spectroscopy, Swinburne University of Technology, Hawthorn, Australia

The adhesion of bacteria to surfaces is an important biological process, but one that has resistedsimple categorization due to the number and complexity of parameters involved. The roughnessof the substrate is known to play a significant role in the attachment process, particularly when thesurface irregularities are comparable to the size of the bacteria and can provide shelter from un-favorable environmental factors. According to this scenario, roughness on a scale much smallerthan the bacteria would not be expected to influence the initial attachment. To test this hypothe-sis, the impact of nanometer-scale roughness on bacterial attachment has been investigated us-ing as-received and chemically etched glass surfaces. The surface modification by etching result-ed in a 70% reduction in the nanoscale roughness of the glass surface with no significant alter-ation of its chemical composition or charge. Nevertheless, the number of bacteria adhering to theetched surface was observed to increase by a factor of three. The increase in attachment was alsoassociated with an alteration in cellular metabolic activity as demonstrated by changes in charac-teristic cell morphologies and increased production of extracellular polymeric substances. The re-sults indicate that bacteria may be more sensitive to nanoscale surface roughness than was pre-viously believed.

Keywords: Bacterial attachment · Glass · Nano-scale roughness

Correspondence: Dr. Elena P. Ivanova, Faculty of Life and Social Sciences,Swinburne University of Technology, PO Box 218, Hawthorn 3122,AustraliaFax: +61-3-9214-5921e-mail: [email protected]

Abbreviations: AFM, atomic force microscopy/microscope; at%, atomicpercent, BHF, buffered hydrofluoric acid; CSLM, confocal scanning laser mi-croscopy/microscope; EPM, electrophoretic mobility; EPS, extracellularpolymeric substances; XPS, X-ray photoelectron spectrometry; XRF, X-rayfluorescence spectroscopy

Received 28 November 2007Revised 28 November 2007Accepted 20 December 2007

© 2008 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim 537

havior. Consequently, a comprehensive theory thatcan reliably predict bacterial attachment behaviorbased on the bacterial surface properties hasproved elusive, despite a considerable amount ofresearch on this topic.

Interest in biofilm formation has been driven bythe importance of bacterial activity in many tech-nological applications, together with a growingability to exert control over surface properties. Inparticular, the ongoing effort to design antibacteri-al surfaces has served to identify a number of sub-stratum surface characteristics that might influ-ence bacterial attachment, including morphology,composition, roughness and porosity [9, 10, 24–29].According to one common scenario of biofilm for-mation, bacterial adhesion is initiated in surface ir-regularities that serve as microenvironmentswhere bacteria are sheltered from unfavorable en-vironmental factors. Bacteria attach to these sites topromote their survival [23, 24, 30, 31].Although theeffects of surface roughness have been studiedover a wide range of physical scales [19, 32–35], toour knowledge it has never been shown that sur-face roughness on a scale much smaller than thebacterium might be a major driver in the initialcourse of bacterial attachment.

This paper presents an investigation of the im-pact of nanometer-scale surface roughness on theattachment strategy of Pseudoalteromonas is-sachenkonii KMM 3549T exposed to glass surfaces.The ‘as-received’ glass surface was modified bychemical etching, as described in Section 2.1.Pseudoalteromonas issachenkonii KMM 3549T isderived from a symbiotrophic association with thedegraded thallus of brown algae Fucus evanescensand has been identified as a novel species of the γ-proteobacteria [36]. Pseudoalteromonas spp. aresoil counterparts of Pseudomonas spp. and repre-sent an abundant group of marine prokaryotes thatcarry out several crucial ecological functions, in-cluding the reduction and/or oxidation of inorgan-ic compounds and the biodegradation of hydrocar-bons and other compounds [37].

2 Materials and methods

2.1 Slide preparation

For the purpose of probing the effects of nanoscaleroughness on bacterial adhesion the surfaces ofstandard glass microscope slides (7105-PPA pre-mium glass slides, Livingstone International) weremodified (etched) by treatment with a buffered so-lution of hydrofluoric acid (BHF) to achieve ananometer-scale variation in surface roughness. In

particular, one half of each slide was immersed inthe BHF etching solution for 20 min [38]. The BHFsolution consisted of 6 parts of 40% ammonium flu-oride (NH4F, Sigma), 1 part of 49% hydrofluoric acid(HF, Asia Pacific Specialty Chemicals) and 14 partsof 36.8% hydrochloric acid (HCl, Merck). All slideswere thoroughly washed with deionized water andstored in 96% alcohol (Aldrich). This procedure al-lowed us to achieve a modification of glass surfacetopography on one half of the glass slide and leavethe other part unchanged. Prior to each experimentthe slides were washed again with deionized waterand placed in sterile petri dishes.

2.2 X-ray fluorescence spectroscopy

Samples of both ‘as-received’ and etched glasseswere prepared by weighing approximately 500 ±0.1 mg of each material into 95% Pt/Au crucibleswith approximately 5 ± 0.1 mg 12–22 lithium tetra-borate/metaborate flux previously dried at 550°C.The sample was fused into a homogenous melt overan oxy-propane flame at temperature of approxi-mately 1055°C for 10 min.A commercially availableiodine-doped cellulose tablet was added approxi-mately 100 s before the molten glass was pouredinto a 32-mm diameter 95% Pt/Au mould heated toa similar temperature.Air jets then cooled the moldand melt for approximately 300 s. The resultingglass discs were analyzed on a Philips PW2404Wavelength Dispersive X-ray fluorescence spec-trometer (XPF) using an in-house calibration andalgorithms developed in the CSIRO-Minerals labo-ratory and control program developed by Philips.

2.3 X-ray photoelectron spectrometry

X-ray photoelectron spectrometry (XPS) was per-formed on an Axis Ultra spectrometer (Kratos An-alytical Ltd., UK), equipped with a monochromaticX-ray source (Al Kα, hν =1486.6 eV) operating at150 W.The spectrometer energy scale was calibrat-ed using the Au 4f7/2 photoelectron peak at bindingenergy (EB =83.98 eV). Photoelectrons emitted at90° to the surface from an area of 700 × 300 μm2

were analyzed with 160 eV survey spectra and a20-eV range was used for high resolution regionspectra for selected elements (O 1s, C 1s, Ca 2p, N1s, Si 2p). The relative atomic concentration of ele-ments detected by XPS was quantified from thearea of peaks in the survey spectra using sensiti-vity factors appropriate for the Kratos instrument.Peaks in the high-resolution region spectra werefitted with synthetic Gaussian-Lorentzian compo-nents after removal of a linear background (usingthe Kratos Vision II software).

Biotechnol. J. 2008, 3, 536–544 www.biotechnology-journal.com

BiotechnologyJournal Biotechnol. J. 2008, 3, 536–544

538 © 2008 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim

2.4 Contact angle measurements

For the glass surfaces, advancing contact angles forwater were measured with the FTA200 (First TenÅngstroms Inc.) using the embedded needlemethod [39, 40]. The droplet contact angle was ob-served with a CCD camera. Measurements wereconducted five times on each surface and the re-sulting movies each delivered up to 100 images foranalysis. After magnification of the image, the con-tact angle was measured using the instrument soft-ware.

Cell surface hydrophobicity was evaluated fromcontact angles measurements on lawns of bacteriausing the sessile drop method. Bacterial cells(OD600 nm = 0.4) in a PBS buffer were deposited oncellulose acetate membrane filters (Sartorius, 0.2μm). The wet filters were air dried at ambient tem-perature (ca. 20ºC) for approximately 30–40 minuntil a “plateau state” was reached. Water dropletswere deposited on each surface. The drop was al-lowed to settle for 2 s without needle contact (forstatic contact angle measurements), images weredigitally saved and contact angle values were ob-tained by processing the image with the instrumentsoftware [41–43].

2.5 Scanning probe microscopy

Atomic force microscopy (AFM) was performed us-ing a Solver P7LS, NT-MDT microscope to imagethe surface morphology and to quantitativelymeasure and analyze the glass surface roughnesson the nanometer scale. Scanning was performedin the semi-contact mode.This reduces the interac-tion between tip and sample and thus allows thedestructive action of lateral forces that exist in con-tact mode to be avoided. In this way the surface fea-tures of the samples were measured with a resolu-tion of a fraction of nanometer, and the surfaceroughness of the investigated areas could be statis-tically analyzed using the standard instrumentsoftware (LS7-SPM v.8.58).

2.6 Cell surface charge

Bacterial zeta potentials provide an indication ofthe overall net surface charge and were obtainedby measuring the electrophoretic mobility (EPM)[28, 44]. The EPM was measured as a function ofionic strength by microelectrophoresis using a zetapotential analyzer (ZetaPALS, Brookhaven Instru-ments Corp).The data were processed with the ac-companying software, which employs the Smolu-chowski equation. Bacterial cell suspension wasprepared as described elsewhere [28, 44]. After the

final wash, the cell pallets were re-suspended in10 mM KCl solution to OD600 nm =1, as suggested byDe Kerchove et al. [28]. This cell solution was thendiluted 1000-fold in 5 mL 10 mM KCl for use for theEPM measurements. Measurements were conduct-ed in an electric field of 2.5 V/cm and frequency of2 Hz. All measurements were done in triplicatesand for each sample the final EPM represents theaverage of five successive ZetaPALS readings, eachof which consisted of 14 cycles per run.

2.7 Bacterial attachment

Prior to each experiment, a fresh bacterial suspen-sion of OD600 nm =0.2 was prepared from P. is-sachenkonii cells grown in marine broth (Difco) atroom temperature (~22°C) for 24 h. A portion of 3–5 mL of bacterial suspension was poured intosterile petri dishes where the glass slides (one glassslide per petri dish) were completely immersed andleft for 12 h of incubation at room temperature(~22°C). After incubation all of the slides werewashed with deionized water and left to dry. Thisapproach allowed the experiments for bacterial at-tachment to be performed under identical condi-tions for each half of each microscope slide. ForAFM, all slides were imaged only after a couple ofhours drying at ambient temperature, which pre-vented the cells from completely drying out.

2.8 Scanning electron microscopy

A FeSEM–ZEISS SUPRA 40VP was used to obtainhigh-resolution images of the bacterial cells. Pri-mary beam energies of 3–15 kV were used, whichallowed features on the sample surface or within afew micrometers of the surface to be observed, re-spectively. For scanning electron microscopy, justbefore imaging, all slides were coated in DynavacCS300 coating unit with carbon and gold to achieveconductivity of the specimen prior to imaging.

2.9 Confocal scanning laser microscopy

Exopolymeric substances produced by the cellsduring the attachment on both surfaces were la-beled with concanavalin A 488 dye (MolecularProbes Inc.). Concanavalin A stock solution wasprepared by dissolving 5 mg in 5 mL 0.1 M sodiumbicarbonate at pH 8.3 and storing at 20°C. Theworking solution was prepared by diluting stocksolution by 1:20 in the same buffer to avoid changesin pH. Glass slides with two regions of differentroughness were prepared as described above.Afterincubation for 23 h at room temperature (~22°C),concanavalin A 488 was added in a ratio of one part

© 2008 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim 539

dye to five parts sample.The slides were then incu-bated for a further period of 1 h under the same in-cubation conditions to allow diffusion. After incu-bation, the samples were washed with sterilizednanopure water and carefully transferred to be ob-served at room temperature (~22°C). The confocalscanning laser microscope (CSLM) Olympus Flu-oview FV1000 Spectroscopic Confocal System wasused. The system includes an inverted microscopeOLYMPUS IX81 [with 20×, 40× (oil), 100× (oil) UISobjectives] and operates with multiple Ar, He andNe laser lines (458, 488, 515, 543, 633 nm).The sys-tem was equipped with a transmitted light differ-ential interference contrast attachment and a CCDcamera (CoolView FDI). Excitation and emissionwavelengths for concanavalin A are 495 and 519nm, respectively.

3 Results and discussion

3.1 Glass surface physico-chemical characteristics

Given that there are a number of factors that caninfluence the initial bacterial attachment, the char-acteristics of both as-received and modified glasssurfaces were investigated in detail. The bulkchemical composition of the glass slides was ob-tained by X-ray fluorescence spectroscopy (XRF).This showed a typical soda-lime glass composition,with the most abundant chemical components inboth samples being SiO2, Na2O, CaO, MgO andAl2O3. The XRF results also indicated that the per-centage of all detected components (20 in total) inboth glass structures was almost identical, with theexception of fluorine, which was found to be pres-ent at a level of 0.37 atomic percent (at%) in themodified glass, compared to 0.24 atomic percent (at%) in the as-received material.This is consistentwith the fact that HF was the main component ofthe etching solution.

XPS results indicate that the most abundant el-ements on both surfaces were O, Si and C.They also

show a modest increase in the relative concentra-tion of O, Si, Ca and Na on the etched surface, whichis consistent with the removal of superficial carbonduring the etching. The levels of Al, F and Fe wereclose to the sensitivity limit for these elements andtherefore the differences in relative concentrationcould not be regarded as significant. The results ofhigh-resolution region spectra shown in Table 1 in-dicate that Si and O were, as expected, predomi-nantly present as silica and Ca was present in the2+ oxidation state.

Analysis of the C1s high-resolution spectra wasconsistent with the presence of hydrocarbons (C-C,C-H), carbon singly bonded to oxygen or nitrogen(C-O, C-N), carbon doubly bonded to oxygen (C=O)and carbonate species (CO3). Although less C wasdetected on the surface of the etched glass com-pared to native glass, which may indicate removalof organic contaminants, the relative concentrationof the various organic species measured by XPS issimilar to that of the native glass. From these re-sults it can be concluded that the etching processdid not cause significant changes in the chemicalcomposition of the glass surface.

The hydrophilicity of the as-received and mod-ified glass surfaces was evaluated via advancingcontact angle measurements. The values listed inTable 2 represent the average of multiple measure-ments. The results indicate an insignificant differ-ence in the surface wettability, which might havebeen caused by the etching solution.

Since no significant differences in chemicalcomposition and wettability could be detected be-tween the two surfaces, further analysis of the sur-face roughness was undertaken by means of scan-ning probe microscopy (SPM). According to thetypical topographic images shown in Fig. 1, themodified glass surface appears uniformlysmoother and without the relatively prominenthigh protrusions of the as-received sample.The re-sultant statistical measures of surface roughnessare listed in Table 2.

Biotechnol. J. 2008, 3, 536–544 www.biotechnology-journal.com

Table 1. Relative contributions of different chemical states assigned to the XPS peaks

Element/transition Assignment As-received glass Etched glassBinding energy (eV) Relative Contrib. Binding energy (eV) Relative Contrib.

N1s C-N 400.5 100 400.3 100

Ca2p Ca2+ 347.9 100 347.5 100

C1s C-C, C-H 285.0 80 285.0 80C-O, C-N 286.5 12 286.5 12

CO3 289.3 4 289.3 4C=O 288.1 3 288.3 4

Si2p SiO2 103.3 100 103.3 100

BiotechnologyJournal Biotechnol. J. 2008, 3, 536–544

540 © 2008 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim

The three conventional roughness parameters,the average surface roughness (Ra), the root meansquare roughness (Rq) and the mean peak to valleyprofile roughness (Rmax), were all approximately70% higher on the native glass surface. However,these parameters do not necessarily provide a sat-isfactory indication of the topographical differ-ences, given the number of relatively prominentprotrusions that are visible in Fig. 1A but not in Fig.1B. The separation between the protrusions on thenative glass is approximately 0.5–1 µm and the dis-tance from the lowest point to the highest can reachup to 16 nm.

An alternative roughness measure that hasbeen suggested for use in the context of biofoulingis the ten-point average roughness Rz, which is de-fined as the difference in height between the aver-age of the five highest peaks and the five lowestvalleys along a profile [32]. In the current context,this parameter has been modified to provide theten-point average roughness over the assessedarea. On this measure, the as-received surface is250% rougher than the etched surface. Overall, allfour parameters (Ra, Rq, Rmax, Rz) suggest that theas-received glass surface was significantly rougherthan the etched glass.

Therefore, contrary to the XRF, XPS and contactangle measurements, the SPM analysis clearly in-dicates that etching has served to modify the topo-

graphical properties of the glass surface. In the ab-sence of any other apparent significant differencesbetween the two surfaces, it would appear that thisstriking nanoscale topographical change is themain point of distinction between them.

3.2 Bacterial surface characteristics

Apart from the substratum surface properties, thehydrophobicity, surface charge and EPS productionof the P. issachenkonii cells have also been charac-terized. It was found that the bacterial cell surfacewas slightly hydrophobic in the range of θ = 64°.The value is similar to those reported for othermembers of the γ-proteobacteria [45]. Cell surfacecharge was inferred from the zeta potential meas-urements.The zeta potential (10 mM) of the bacte-rial cells was estimated as –34 ± 3 mV. As previous-ly mentioned, both surface charge and wettabilitycharacteristics are extremely dependent on theproduction of extracellular substances and canvary significantly between bacteria belonging tothe same generic species. A surface wettability ofθ = 64° and surface charge of –34 ± 3 mV indicatethat the surface characteristics of P. issachenkoniicells are similar to those of other bacteria belong-ing to the γ-protobacteria [23, 30, 34]. They also in-dicate that P. issachenkonii cells may not have astrong affinity to the hydrophilic glass surface.

Table 2. Surface wettability and roughness parameters for the as-received and etched glass

Sample Contact angle Surface roughness (nm)Water Ra Rq Rmax Rz

As-received glass 44.8 ± 0.1° 2.1 2.8 27.8 12.2Etched glass 41.6 ± 0.1° 1.3 1.6 16.2 4.8

Figure 1. AFM images of as-received (A) and etched (B) glass surfaces. It is evident that the treatment with BHF has resulted in a significant reduction ofsurface roughness on the nanometer scale. The imaged areas are about 5 × 5 μm2 and 5 × 6 μm2, respectively.

14nm 14nm

© 2008 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim 541

3.3 Bacterial attachment

After determining the basic physico-chemicalcharacteristics of both bacteria and substrata, theattachment behavior of P. issachenkonii was ob-served on both the as-received and modified glasssurfaces.The particular morphology of the bacteriaafter adhering to the surfaces was examined bymeans of high-resolution scanning electron micro-scope, as shown by the images in Fig. 2.

An initial inspection of the bacterial attachmentrevealed striking differences in the bacterial re-sponse to the two surface regions. The number ofattached cells observed in the scanning electronmicroscope images (1000× magnification) wastransformed into a number of bacteria per unit areaand was tentatively estimated to be 36 000 cells/mm2 on the as-received glass surface, while thebacterial density increased by a factor of three onthe modified glass surface, reaching approximately

110 000 cells/mm2. These densities have estimatederrors of approximately 10% due to local variabilityin the coverage.

In addition to the obvious numerical differ-ences, clear changes were observed in the cell mor-phology and the production of extracellular poly-meric material (presumably extracellular polysac-charides). Morphological differences in the cells at-tached to both surfaces after 12-h incubation wereseen in high magnification scanning electron mi-croscope images (inserts of Fig. 2) and were con-firmed by AFM, as shown in Fig. 3 and Table 3.

The different size of the bacterial cells can bequantitatively compared on the basis of Figs. 3Aand B. According to the original description [36],cells of P. issachenkonii are 0.7–0.9 µm wide and1.0–1.2 µm long. In these experiments, the majorityof the cells attached to the as-received glass were2 µm long, 1 µm wide and 140 nm high (Fig. 3A andTable 3). However, on the modified glass surfaces,

Biotechnol. J. 2008, 3, 536–544 www.biotechnology-journal.com

Figure 3. AFM images demonstrating cell morphology on the as-received (A) and the modified (B) glass surfaces after 12-h incubation. Some granular EPSparticles are observed on the modified glass surface (Ba, Bb). The imaged areas are about 5 × 5 μm2 in each case.

Figure 2. Representative scanning electron microscopy images of attached bacteria, taken from non-treated (A) and modified (B) glass surfaces after 12-hincubation. Both images (1000× magnification) provide a general overview of the bacterial attachment and illustrate the different attachment patterns.Inserts in the upper left corners (5000× magnification) reveal a morphological transformation in the cells attached to the modified glass surface. The mor-phological transformation is shown by increases in cell size and production of extracellular polymeric substances. Scale bars on both images: 10 μm.

160nm 250nm

BiotechnologyJournal Biotechnol. J. 2008, 3, 536–544

542 © 2008 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim

the average width of the bacterial cells was foundto be 1.3 µm, the length 2.9 µm and the averageheight 170 nm (Fig. 3B and Table 3). These differ-ences in cell morphology may result from differentstrategies used by the P. issachenkonii cells to at-tach to the glass surfaces. It is suggested that thedifference in surface roughness provoked the pro-duction of different quantities of EPS, which led tochanges in cell morphology. In general, cells on theas-received glass surface appeared smaller andflatter than cells on the modified glass surface.Apart from the EPS coating the cells, additionalquantities of EPS [80–120 nm in height at points (a)and (b) in Fig. 3B] were also found on the modifiedglass surface.This is indicative of the surface mod-ification strategy utilized by P. issachenkonii to bet-ter sustain their existence on this surface.

The production of extracellular substances dur-ing the process of adhesion was observed usingCSLM.The attachment pattern of bacterial cells ontwo regions of glass surfaces after 24 h of incuba-tion is presented in Fig. 4. The number of bacterialcells remained greater on the modified glass sur-faces after this extended incubation period. It ap-pears that the cells attached to the modified glass

surfaces began to form a multilayer structure andproduced greater quantities of EPS, according tothe fluorescence images.

As concanavalin A specifically binds to α-mannopyranosyl and α-glucopyranosyl residues, itcan be assumed that these sugars are componentsof EPS found on the cell surfaces. However, thegranular EPS observed by AFM on the etched glasssurface [see points (a) and (b) in Fig. 2B] was notdetected in the confocal images, suggesting a dis-tinct chemical composition for this type of EPS.This is an indication that changes in the nanoscalesurface roughness might induce the P. issachen-konii cells to produce different types of EPS duringthe attachment process. It has repeatedly been re-ported that bacteria produce different EPS de-pending on the environmental conditions (e.g.,[46]), but we are not aware of nanoscale surface to-pography being implicated in this process before.However, identification of the chemical composi-tion of the EPS produced by P. issachenkonii cellson both types of glass surface remains a challeng-ing task due to the small amounts of material avail-able for analysis.

4 Concluding remarks

The results presented here suggest that nanoscalesurface roughness may exert a greater influence onbacterial adhesion than was previously believed,and should therefore be considered as a parameterof primary interest alongside other well-recog-nized factors that control initial bacterial attach-ment. This investigation has shown that, under

Table 3. Bacterial cell dimensions after 12-h incubation on both surfaces,as inferred from AFM measurements

Dimension (μm) On the as-received On the modified glass surface glass surface

Length 2.1 2.9Width 1.0 1.3Height 0.14 0.17

Figure 4. CLSM images of EPS (stained green with concanavalin A 488) overlaying conventional images of cells attached to the as-received (A) and modi-fied (B) glasses. The scale bar is 10 μm.

© 2008 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim 543

similar conditions, bacterial adhesion to glass wassignificantly influenced by nanometer scalechanges in the surface roughness and topography.The bacterial response was translated into a re-markable change in cellular metabolic activity asdemonstrated by the characteristic cell morpholo-gies and production of extracellular polymeric sub-stances. The suggestion that bacteria may be sus-ceptible to nanoscale surface roughness casts seri-ous doubt on the conventional wisdom thatsmoother surfaces represent a more repellent en-vironment to bacteria. The effect of nanoscale sur-face roughness on bacterial adhesion has impor-tant implications for designing surfaces for use insurgical implants, the food industry and sterile en-vironments such as hospitals and pharmaceuticallaboratories.

We thank Hans Brinkies, Swinburne University ofTechnology, Faculty of Engineering and IndustrialSciences, for his assistance with the scanning elec-tron microscopy; Daniel White, Swinburne Universi-ty of Technology, Centre for Atom Optics and Ultra-fast Spectroscopy for his assistance in etching theglass surfaces, Grant van Reissen, LaTrobe Univer-sity, for the XPS analysis, and Steve Peacock, CSIROMinerals, for the XRF analysis.

The authors have declared no conflict of interest.

5 References

[1] Beech, B. I., Sunner, J., Biocorrosion: Towards understand-ing interactions between biofilms and metals. Curr. Opin.Biotechnol. 2004, 15, 181–186.

[2] Parsek, M. R., Singh, P. K., Bacterial biofilms: An emerginglink to disease pathogenesis. Annu. Rev. Microbiol. 2003, 57,677–701.

[3] Simoni, S. F., Bosma, T. N. P., Harms, H., Zehnder, A. J. B., Bi-valent cations increase both the subpopulation of adheringbacteria and their adhesion efficiency in sand columns. En-viron. Sci. Technol. 2000, 34, 1011–1017.

[4] Teixeira, P., Oliveira, R., Influence of surface characteristicson the adhesion of Alcaligenes denitrificans to polymericsubstrates. J. Adhes. Sci. Technol. 1999, 13, 1243–362.

[5] Benito, Y., Pin, C., Luisa Martin, M., Luisa Garcia, M. et al.,Cell surface hydrophobicity and attachment of pathogenicand spoilage bacteria to meat surfaces. Meat Sci. 1997, 45,419–425.

[6] Whittaker, C. J., Klier, C. M., Kolenbrander, P. E., Mechanismsof adhesion by oral bacteria. Annu. Rev. Microbiol. 1996, 50,513–552.

[7] Fletcher, M., Loeb, G. I., Influence of substratum character-istics on the attachment of a marine pseudomonad to solidsurfaces. Appl. Environ. Microbiol. 1979, 37, 67–72.

[8] Costerton, J.W., Lewandowski, Z., DeBeer, D., Caldwell, D. etal., Biofilms, the customized microniche. Biophys. J. 1994,176, 2137–2142.

[9] Taylor, R. L., Verran, J., Lees, G. C., Ward, A. J. P., The influ-ence of substratum topography on bacterial adhesion topolymethyl methacrylate. J. Mater. Sci. 1998, 9, 17–22.

[10] Verran, J., Drucker, D. B.,Taylor, C. J., Feasibility of using au-tomatic image-analysis for measuring deposition of Strep-tococcus mutans onto glass, in terms of percentage coverageand mean clump size. Microbios 1980, 29, 161–169.

[11] Pereira, M.A.,Alves, M. M.,Azeredo, J., Mota, M., Oliveira, R.,Influence of physico-chemical properties of porous micro-carriers on the adhesion of an anaerobic consortium. J. Ind.Microbiol. Biotechnol. 2000, V24, 181–186.

[12] Castellanos, T., Ascencio, F., Bashan, Y., Cell-surface hy-drophobicity and cell-surface charge of Azospirillum spp.FEMS Microbiol. Ecol. 1997, 24, 159–172.

[13] Bos, R., van der Mei, H. C., Busscher, H. J., Physico-chemistryof initial microbial adhesive interactions – Its mechanismsand methods for study. FEMS Microbiol. Rev. 1999, 23,179–230.

[14] Busscher, H. J., van der Mei, H. C., Physico-chemical inter-actions in initial microbial adhesion and relevance forbiofilm formation. Adv. Dent. Res. 1997, 11, 24–32.

[15] Cao, T., Tang, H., Liang, X., Wang, A. et al., Nanoscale inves-tigation on adhesion of E. coli to surface modified siliconeusing atomic force microscopy. Biotechnol. Bioeng. 2006, 94,167–176.

[16] Pringle, J. H., Fletcher, M., Influence of substratum hydra-tion and adsorbed macromolecules on bacterial attachmentto surfaces. Appl. Environ. Microbiol. 1986, 51, 1321–1325.

[17] Eboigbodin, E. K., Newton, A. R. J., Routh, F. A., Biggs, A. C.,Role of non-adsorbing polymers in bacterial aggregation.Langmuir 2005, 21, 12315–12319.

[18] Wong, H.-C., Chung, Y.-C. ,Yu, J.-A., Attachment and inacti-vation of Vibrio parahaemolyticus on stainless steel andglass surface. Food Microbiol. 2002, 19, 341–350.

[19] Sharon, N., Carbohydrates as future anti-adhesion drugs forinfectious diseases. Biochim. Biophys. Acta 2006, 1760,527–537.

[20] Stoodley, P., Sauer, K., Davies, D. G., Costerton, J.W., Biofilmsas complex differentiated communities. Annu. Rev. Microbi-ol. 2002, 56, 187–209.

[21] Bell, C. H., Arora, B. S., Camesano, T. A., Adhesion ofPseudomonas putida KT2442 is mediated by surface poly-mers at the nano- and microscale. Environ. Eng. Sci. 2005, 22,629–641.

[22] Fletcher, M., Bacterial Attachment in Aquatic Environments:A Diversity of Surface and Adhesion Strategies, Wiley-Liss,New York 1996.

[23] Li, B., Logan, B. E., Bacterial adhesion to glass and metal-ox-ide surfaces. Colloids Surfaces B: Biointerfaces 2004, 36,81–90.

[24] Shellenberger, K., Logan, B. E., Effect of molecular scaleroughness of glass beads on colloidal and bacterial deposi-tion. Environ. Sci. Technol. 2002, 36, 184–189.

[25] Hazan, Z., Zumeris, J., Jacob, H., Raskin, H., et al., Effectiveprevention of microbial biofilm formation on medical de-vices by low-energy surface acoustic waves. Antimicrob.Agents Chemother. 2006, 50, 4144–4152.

[26] Riedewald, F., Bacterial adhesion to surfaces:The influenceof surface roughness. PDA J. Pharm. Sci. Technol. 2006, 60,164–171.

[27] Satriano, C., Messina, G. M. L., Carnazza, S., Guglielmino, S.,Marletta, G., Bacterial adhesion onto nanopatterned poly-mer surfaces. Mater. Sci. Eng. C. 2006, 26, 942–946.

Biotechnol. J. 2008, 3, 536–544 www.biotechnology-journal.com

BiotechnologyJournal Biotechnol. J. 2008, 3, 536–544

544 © 2008 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim

[28] de Kerchove, A. J., Elimelech, M., Relevance of electrokinet-ic theory for “soft” particles to bacterial cells: implication forbacterial adhesion. Langmuir 2005, 21, 6462–6472.

[29] Shimp, R. J., Pfaender, F. K., Effects of surface area and flowrate on marine bacterial growth in activated carboncolumns. Appl. Environ. Microbiol. 1982, 44, 471–477.

[30] Jones, J. F.,Velegol, D., Laser trap studies of end-on E. coli ad-hesion to glass. Colloids Surfaces B: Biointerfaces 2006, 50,66–71.

[31] Chae, M. S., Schraft, H., Truelstrup Hansen, L., Mackereth,R., Effects of physicochemical surface characteristics of Lis-teria monocytogenes strains on attachment to glass. Food Mi-crobiol. 2006, 23, 250–259.

[32] Whitehead, A. K., Verran, J., The effect of surface topogra-phy on the retention of microorganisms. Food Bioprod.Process. 2006, 84, 253–259.

[33] Emerson, R. J., Bergstrom, T. S., Liu, Y., Soto, E. R. et al., Mi-croscale correlation between surface chemistry, texture, andthe adhesive strength of Staphylococcus epidermidis. Lang-muir 2006, 22, 11311–11321.

[34] Li, B., Logan, B. E., The impact of ultraviolet light on bacte-rial adhesion to glass and metal oxide-coated surface. Col-loids Surfaces B: Biointerfaces 2005, 41, 153–161.

[35] Bruinsma, G. M., Rustema-Abbing, M., van der Mei, H. C.,Busscher, H. J., Effects of cell surface damage on surfaceproperties and adhesion of Pseudomonas aeruginosa. J. Mi-crobiol. Methods 2001, 45, 95–101.

[36] Ivanova, E. P., Sawabe,T.,Alexeeva,Y.V., Lysenko,A. M. et al.,Pseudoalteromonas issachenkonii sp. nov., a bacterium thatdegrades the thallus of the brown alga Fucus evanescens.Int. J. Syst. Evol. Microbiol. 2002, 52, 229–234.

[37] Garrity, G. M., Bergey’s Manual of Systematic Bacteriology,2nd edn, Williams & Wilkins, Baltimore 1984.

[38] White, J. D., Stoddart, R. P., Nanostructured optical fiber withsurface-enhanced Raman scattering functionality. OpticsLett. 2005, 30, 598–600.

[39] Öner, D., McCarthy, T. J., Ultrahydrophobic surfaces. Effectsof topography length scales on wettability. Langmuir 2000,16, 7777–7782.

[40] Quéré, D., Lafuma, A., Bico, J., Slippy and sticky microtex-tured solids. Nanotechnology 2003, 14, 1109–1112.

[41] Dong, H., Onstotta, T. C., Kob, C.-H. A., Hollingsworth, A. D.,Brown, D. G., Mailloux, B. J., Theoretical prediction of colli-sion efficiency between adhesion-deficient bacteria andsediment grain surface. Colloids Surfaces B, Biointerfaces2002, 24, 229.

[42] Korenevsky, A., Beveridge, T. J., The surface physicochem-istry and adhesiveness of Shewanella are affected by theirsurface polysaccharides. Microbiology 2007, 153, 1872–1883.

[43] Bakker, D. P., Busscher, H. J., van der Mei, H. C., Bacterialdeposition in a parallel plate and a stagnation point flowchamber: Microbial adhesion mechanisms depend on themass transport conditions. Microbiology 2002, 148, 597–603.

[44] Eboigbodin, E. K., Newton, A. R. J., Routh, F. A., Biggs, A. C.,Bacterial quorum sensing and cell surface electrokineticproperties. Appl. Microbiol. Biotechnol. 2006, 73, 669–675.

[45] van der Mei, H. C., Bos, R., Busscher, H. J., A reference guideto microbial cell surface hydrophobicity based on contactangles. Colloids Surfaces B: Biointerfaces 1998, 11, 213–221.

[46] de Rezende, C. E.,Anriany,Y., Carr, L. E., Joseph, S.W.,Wein-er, R. M., Capsular polysaccharide surrounds smooth andrugose types of Salmonella enterica serovar TyphimuriumDT104. Appl. Environ. Microbiol. 2005, 71, 7345–7351.