The effect of surface properties on the strength of attachment of fungal spores using AFM perpendicular force measurements

Colloids and Surfaces B: Biointerfaces 82 (2011) 483–489

Contents lists available at ScienceDirect

Colloids and Surfaces B: Biointerfaces journal homepage: www.elsevier.com/locate/colsurfb

The effect of surface properties on the strength of attachment of fungal spores using AFM perpendicular force measurements Kathryn A. Whitehead a,∗ , Ted Deisenroth b , Andrea Preuss c,1 , Christopher M. Liauw a , Joanna Verran a a b c

Manchester Metropolitan University, Manchester, UK Ciba Speciality Chemicals Corporation, Tarrytown, New York, USA Ciba Spezialitatenchemie, Grenzach, Germany

a r t i c l e

i n f o

Article history: Received 2 April 2010 Received in revised form 2 August 2010 Accepted 4 October 2010 Available online 8 October 2010 Keywords: Fungi Polymeric surfaces Atomic force microscopy (AFM) Hydrophobicity Force

a b s t r a c t Polymeric substrata may be biodegraded by fungal species resulting in damaged, weakened and unsightly materials. This process typically begins with fungal spore attachment to the surface. In order to better understand the processes that precedes a biofouling event, fungal spore attachment to a range of surfaces, was determined using perpendicular force measurements. This was carried out using atomic force microscope cantilevers modified with fungal spores from Aspergillus niger 1957 (5 ␮m diameter, non-wettable, spherical), Aspergillus niger 1988 (5 ␮m diameter non-wettable, spikey) or Aureobasidium pullulans (5 ␮m–10 ␮m sized, wettable, ellipsoidal). The strength of attachment of the spores was determined in combination with seven surfaces (nitric acid cleaned glass, cast poly(methylmethacrylate) sheet [c-PMMA], polytetrafluoroethylene [PTFE], silicon wafers spin coated with poly(3-methacryloxypropyltrimethoxy silane (␥-MPS)-co-methylmethacrylate (MMA)) [p(␥-MPSco-MMA)], poly (␥-MPS-co-lauryl methacrylate) [p(␥-MPS-co-LMA)] [both in a ratio of 10–90], PMMA dissolved in a solvent [PMMAsc] and silicon wafers). Perpendicular force measurements could not be related to the Ra values of the surfaces, but surface wettability was shown to have an effect. All three spore types interacted comparably with the surfaces. All spores attached strongly to c-PMMA and glass (wettable surfaces), and weakly to PTFE, (p(␥- MPS-co-LMA)) (non-wettable) and (p(␥-MPS-co-MMA)). Spore shape also affected the strength of attachment. Aureobasidium pullulans spores attached with the widest range of forces whilst A. niger 1957 attached with the smallest. Findings will inform the selection of surfaces for use in environments where biofouling is an important consideration. © 2010 Elsevier B.V. All rights reserved.

1. Introduction Biofouling is a common problem faced by systems as diverse as marine surfaces [1,2], interior and exterior surfaces and paint [3], stone [4], water distribution systems [5], books and paper [6], textiles [7], window glass [8], stone monuments [9], cultural heritage [10] and synthetic polymers [11–13]. Coatings developed to reduce biofouling of engineered surfaces do not always perform as expected based on their native properties [14] due to microbial and fungal spore attachment. Spores are propagative, microbial vectors that can survive extremes of chemical and physical attack in harsh environments [15]. Aerial transmission of fungal spores has serious implications in agriculture, medicine and the food industry

∗ Corresponding author at: Manchester Metropolitan University, Department of Biology, Room E240, John Dalton Extension Chester St, Manchester, United Kingdom. Tel.: +44 (0) 161 247 1157; fax: +44 (0) 161 247 6365. E-mail address: [email protected] (K.A. Whitehead). 1 Current address: Product Development, CoE Infection Control, B. Braun Medical AG, Seesatz CH-6204, Sempach, Switzerland. 0927-7765/$ – see front matter © 2010 Elsevier B.V. All rights reserved. doi:10.1016/j.colsurfb.2010.10.001

[16–18], thus an improved understanding of the interactions that affect spore attachment may help in the development of strategies to reduce spore adherence [19]. It is known that the physical structure of the outer surfaces of spores varies considerably [19]. Aspergillus niger spores are unicellular, hydrophobic (non-wettable) and have relatively little chemical surface variation [15], but may vary significantly in shape. Aureobasidium pullulans is ubiquitous in the environment, and colonises many habitats [20]. It is a principal colonizer of painted wood surfaces [21] and was found to be the dominant fungus causing biodeterioration of PVC films during outdoor trials in Florida [22]. The colony is usually black pigmented due to melanin in the cell wall [23]. This material is thought to physically protect the organism and to aid adhesion of the organism to surfaces [24,25]. Attachment of a microorganism to a solid substratum is the first component in a series of events that occurs during the colonization of the surface. Whilst bacterial attachment has been extensively studied [26–29], considerably less attention has been given to fungal attachment with the exception of the opportunistic pathogen Candida albicans [30,31].

484

K.A. Whitehead et al. / Colloids and Surfaces B: Biointerfaces 82 (2011) 483–489

Table 1 Chemical name, Ra and contact angle measurements of surfaces. n = 5 for Ra and contact angle measurements. ± values indicate standard deviation from the mean. Acronym

Chemical name

Ra (nm)

Glass (p(␥-MPS-co-MMA))

Nitric acid cleaned glass Silicon wafers spin coated with poly(3-methacryloxypropyltrimethoxy silane (␥-MPS)-co-methylmethacrylate) [in a ratio of 10–90] Standard cast poly(methylmethacrylate) sheet Polytetrafluoroethylene Silicon wafers spin coated with poly(␥-MPS-co-laurylmethacrylate) [in a ratio of 10–90] PMMA dissolved in a solvent Silicon wafer

117.8 ± 8.2 32.0 ± 8.5

c-PMMA PTFE (p(␥-MPS-co-LMA)) PMMAsc Si

Initial spore attachment is due to physicochemical interactions between the biological surfaces and the substratum [19]. Substratum hydrophobicity has generally been acknowledged to be one of the factors contributing to the ability of microorganisms to adhere to another surface [32]. The method of substratum production and its chemistry influence both surface wettability and roughness. Wear of surfaces through their production process, surface finish, abrasion, cleaning and impact damage also increase surface roughness [33]. This can introduce topographical features which may increase the retention of microorganisms [34–36]. Advances in the use of atomic force microscopes (AFM) have permitted the direct measurement of the force of attachment of a single spore in the direction normal to the interacting surfaces. This is achieved by the immobilisation of a single fungal spore at the end of a cantilever, thus creating a particle probe [37,38]. Force distance curves are recorded for interactions between the particle probes and substrata and using the spring constant and a measurement of the force where the cantilever and the surface are apart (zero of the force), data is converted into a numerical force value. This study aimed to determine the effect of surface properties (chemistry, topography and wettability) on fungal spore attachment. Fungal spores were selected based on differing morphology and wettability characteristics and on the importance of the species in biofouling and biodegradation of environmental surfaces. Fungal spores were used to produce particle probes by attachment to the AFM cantilever, and their strength of attachment to surfaces with defined chemistries, topographies and wettabilities was determined.

3.0 ± 0.3 182.9 ± 13.6 4.1 ± 0.9 2.0 ± 0.3 0.6 ± 0.0

Contact angle (◦ ) 57 ± 5 61 ± 5

71 ± 5 107 ± 5 108 ± 15 62 ± 2 42 ± 2

copolymerised with MMA, PMMAsc was made by spin coating 3-methacryloxypropyltrimethoxysilane (␥-MPS) copolymerised with methyl methylacrylate (MMA) onto a silicon wafer and (p(␥-MPS-CO-LMA)) was lauryl methacrylate copolymerised with ␥-MPS. 2.1.3. Fourier transformation infra red spectroscopy (FTIR) A Thermo-Nicolet Continuum FTIR microscope attached to a Thermo Nicolet Nexus FTIR spectrophotometer bench was used for FTIR analysis of the surfaces. The microscope was fitted with a type A MCT detector and the aperture was set to its maximum size of 200 ␮m × 200 ␮m. The spectra were acquired using Omnic 5.2 software (Nicolet, Madison, USA) and were made up of 32 scans with resolution set to 4 cm−1 . 2.1.4. Surface wettability Surface wettability was determined using contact angle measurements at room temperature. The sessile drop technique was used with 5 ␮l volumes of HPLC grade water (BDH, UK) using a Kruss instrument (KRÜSS GMBH, Hamburg, Germany). 2.1.5. Ra values of the surfaces Substrata images and roughness measurements were obtained using an Explorer AFM (Veeco Instruments, Cambridge, UK). Surface roughness analysis was carried out in contact mode using a cantilever with a spring constant of 0.05 N m−1 or in non-contact mode with a non-contact cantilever with a spring constant of 50 N m−1 (Veeco Instruments, Cambridge, UK).

2. Experimental methods 2.2. Mycology 2.1. Surface production and analysis Seven different surfaces presenting different properties were used in this study (Table 1). 2.1.1. Commercially purchased surfaces Silicon wafers were purchased from Montco Technologies (PA, USA), PTFE from Direct Plastics (Yorkshire, UK) and cast (c-)PMMA sheet from Manchester Plastics (Manchester, UK). Glass cover slips (20 mm × 20 mm) were purchased from Scientific Laboratory Supplies (UK). Glassware was pre-cleaned in 69% fuming nitric acid followed by a solvent wash which consisted of rinsing the glassware for 5 min in each of distilled water, acetone, methanol, ethanol and finally in sterile distilled water. Nitric acid and solvents were obtained from BDH (Basingstoke, UK). 2.1.2. Spin coated surfaces The spin coated surfaces were produced by spin coating silicon wafers with dissolved polymers. The desired polymer (or polymerising mixture) was dropped onto silicon wafer disks so that the disk was covered. Samples were spun at 2000 rpm for 10–15 s. (p(␥-MPS-MMA)) was made with ␥-MPS

2.2.1. Cultivation of fungi from lyophilizates Fungal cultures were a kind gift from Ciba Spezialitatenchemie (Germany). These were re-suspended from freeze dried lyophilizates. The “lyo flasks” were opened using a glass cutter, and the cotton wool was removed with tweezers. The “lyo pill” or pellet was transferred into a test tube and 3 ml NaCl–Triton X-100 (A. niger spp.) or Sabouraud broth (Aureobasidium pullulans) (BDH, Basingstoke, UK) was added. NaCl–Triton X-100 was made by suspending 1 ml of a v/v 1% solution in 100 ml sterile distilled water, into which 0.85 g sodium chloride (Sigma, Dorset, UK) was dissolved. The suspension was vortexed for 5 min or until the pellet was dispersed. The fungal suspension was dispensed onto Sabouraud agar plates (BDH, Basingstoke, UK). The plates were incubated for 3–21 days at 29 ◦ C. Following growth of the fungi, a second transfer was made. A cotton swab was dipped into diluent (NaCl–Triton X-100 for A. niger spp. and Sabouraud broth for Aureobasidium pullulans) and was wiped across the culture, then spread onto a fresh agar plate. Inoculated plates were incubated for 3–28 days at 29 ◦ C. After good growth of the fungi, a third transfer was made. This transfer was used to obtain the “working” cultures from which spores were harvested.

K.A. Whitehead et al. / Colloids and Surfaces B: Biointerfaces 82 (2011) 483–489

485

Cultures for short term storage were made on agar slopes. Slopes were inoculated with a swab dipped into the appropriate diluents. Inoculated slopes were incubated for 3–21 days at 29 ◦ C. Short term storage cultures were stored at 4 ◦ C for up to 12 months for long term storage. Fungal slopes were covered in a layer of sterile mineral oil (Sigma, Poole, UK), and stored at 4 ◦ C.

3 min, at 2500 V, in argon gas at a power of 18–20 mA. Images of substrata were obtained using a JEOL JSM 5600LV scanning electron microscope. EDX chemical analysis of spores was carried out to a 1 ␮m depth using a Link Pentafet detector (Oxford Instruments, Buckinghamshire, UK), with Inca software (Oxford Instruments, Buckinghamshire, UK).

2.2.2. Spore suspensions Fungi were inoculated from a short term or a long term agar slope on agar using a sterile swab dipped in NaCl–Triton X-100 (A. niger spp.) or Sabouraud broth (Aureobasidium pullulans). Inoculated plates were incubated for 3–21 days at 29 ◦ C. Following fungal growth, 5 ml of sterile diluent was pipetted onto the fungal culture. Spores were removed from the culture by rubbing a sterile glass Pasteur pipette gently over the surface of the culture. The suspension was transferred into a sterile beaker with a sterile magnetic stirrer. This process was repeated several times until all the visible sporulating growth was removed from the agar. The suspension was stirred for 30 min then filtered through a funnel filled with glass wool (VWR, Dorset, UK) to obtain a homogeneous spore suspension. Spores were harvested at 3000 × g for 10 min, washed three times in sterile distilled water, and re-suspended to an optical density of 1.0 at 610 nm which equated to 6.53 ± 1.88 × 106 spores cm2 . Spore suspensions were stored at 4 ◦ C and used within two weeks. Spores were checked for purity by light microscopy before each use.

2.2.5. Light microscopy One hundred microlitres of spore suspension was pipetted onto a glass slide and air dried in a class 2 microbiological containment hood. Substrata were visualised using light microscopy (Nikon Eclipse E600, Surrey, UK). The microscope was mounted with an F-View II digital camera (Soft Imaging System Ltd., Helperby, UK, supplied by Olympus, Hertfordshire, UK). This system used a Cell F Image Analysis package (Olympus, Hertfordshire, UK).

2.2.3. Microbial adhesion to hydrocarbons (MATH) assay Spore surface affinity to hydrocarbons was measured according to an adapted method of the MATH assay described by Rosenberg and Doyle [39]. Spore suspensions were washed 3 times in PUM Buffer pH 7.1 (PUM Buffer; 22.2 g l−1 K2 HPO4 ·3H2 O (BDH, Hampshire, UK) 7.26 g l−1 KH2 PO4 (BDH, Basingstoke, UK); 1.8 g l−1 urea (Sigma, Poole, UK); 0.2 g l−1 MgSO4 ·7H2 O (BDH, Basingstoke, UK)) and re-suspended to an OD 1.0 at 520 nm. Five millilitres of washed spores suspended in PUM buffer were added to 15 mm diameter round bottom glass test tubes. One millilitre of n-hexadecane (BDH, Basingstoke, UK) was added to the test suspension. Suspensions were incubated at 29 ◦ C for 10 min to equilibrate. The suspensions were mixed for 2 min, and then incubated for 30 min at 29 ◦ C. The weaker aqueous phase was transferred to cleaned test tubes, and the OD determined at 520 nm. The calculation used to determine spore affinity to hydrocarbons was as that of Rosenberg and Doyle [39]; %Adhesion = 1 −

A A0

× 100

(1)

A0 is the optical density of the spore suspension before mixing. A is the optical density following mixing with hydrocarbon and extraction of the aqueous phase. 2.2.4. Inoculation and preparation of spores for scanning electron microscopy (SEM) and electron dispersive X-ray (EDX) analysis One hundred microlitres of spore suspension was pipetted onto to a 10 mm × 10 mm polished silicon wafer surface (Montco Silicon Technologies, PA, US) and air dried in a class 2 microbiological containment hood. Following this initial drying, surfaces were placed for 1 week in a desiccator containing phosphorous pentoxide (Sigma, Dorset, UK). Substrata plus retained spores were immersed in 4%, v/v gluteraldehyde (Agar Ltd., Essex, UK) for 24 h at 4 ◦ C. After fixing, substrata were washed gently with distilled water. Prior to examination, samples were stored at room temperature, in a desiccator containing phosphorous pentoxide. For SEM imaging, samples were fixed onto stubs for gold sputter coating, which was carried out using a Polaron E5100 (Milton Keynes, UK) SEM sputter coater. Samples were sputter coated at a vacuum of 0.09 mbar, for

2.3. Strength of attachment measurements 2.3.1. Modification of tipless cantilevers to produce particle probe Tipless cantilevers (Veeco, Cambridge, UK) with a spring constant of 0.12 Nm−1 were glued using two phase silver mounting adhesive onto cantilever stubs (Veeco, Cambridge, UK). Ten microlitres of washed spore suspension were pipetted into a new, sterile, clean Petri dish. The diluent was evaporated off in a class 2 flow hood. Samples were thoroughly dried in a phosphorous pentoxide desiccator for three days. A 20 cm × 20 cm glass cover slip was attached to a AFM mounting disc (Veeco, Cambridge, UK) using double sided sticky tape. A small number of dried spores were removed from the Petri dish and placed onto the coverslip. A small amount of cyanoacrylate gel (Bostik, UK) was added to the coverslip. The mounting disc with the attached coverslip was placed on the AFM. Using the AFM camera and XY automated translation stage, the tipless cantilever was moved to the edge of the cyanoacrylate gel, and was lowered in the z plane until the cantilever momentarily touched the gel. The cantilever was quickly moved in the z plane. The cantilever was then moved across the coverslip until a suitable spore was found. The cantilever was again lowered in the z plane until it touched the spore. The cantilever was left in place for 10 s before being lifted vertically. The cantilever was removed from the AFM and left for 24 h to allow the adhesive to fully cure. The quality/validity of the spores glued to the cantilevers was checked using both light and electron microscopy (data not shown). It was found that using repeated control experiments that the spore-cantilevers could each be used for up to 20 force measurements before they became unstable. Results are presented as the mean ± standard deviation of each measured value. 2.3.2. Strength of spore attachment to the substratum Before each experiment the spring constant of the cantilever was determined by measuring the mechanical response of the cantilever to thermal noise as a function of time using the AFM software. To measure the adhesion force between the particle probes and the substratum, the particle probe was brought into momentary contact with the surface. AFM strength of attachment measurements were obtained from force–distance curves. The spring constant of the cantilever, zero of the force and the cantilever deflection (d) were converted into a force (F) using Hooke’s law [37,38] F = −kd

(2)

where k is the cantilever spring constant. The cantilever deflection (distance (d)) was corrected by plotting F as a function of (z–d), where z is the vertical displacement of the piezoelectric scanner. To calculate the force the spring constant was multiplied by the displacement (Hookes Law), and the zero of the force was subtracted from the image setpoint. This value was converted to nN from nA

K.A. Whitehead et al. / Colloids and Surfaces B: Biointerfaces 82 (2011) 483–489

% Adsorbance in polar phase

486

100 80 60 40 20 0 A. niger

A. niger

1957

1988

Aureobasidium pullulans

Fig. 1. MATH results demonstrating percentage adherence to the polar phase of fungal spores. Error bars demonstrate standard deviation of the mean (n = 10).

by multiplying the applied force by the reciprocal of the slope and the cantilever spring constant. The spring constant was individually determined for each cantilever before each experiment.

Fig. 3. Light microscopy of Aureobasidium pullulans demonstrating EPS surrounding spores.

40

3. Results

A. niger 1957

35

A. niger 1988

30

% weight

Only a small percentage of either the A. niger 1957 or A. niger 1988 spores adsorbed into the polar phase during the MATH assay whereas a high number of the Aureobasidium pullulans spores absorbed into the polar phase (Fig. 1). Spores of A. niger 1957 were of 5 ␮m diameter and spherical (Fig. 2a) whereas A. niger 1988 produced 5 ␮m diameter spherical spores with spikey appendages (Fig. 2b). Aureobasidium pullulans produced 5 ␮m–10 ␮m ellipsoidal spores (Fig. 2c). Aureobasidium pullulans excreted large amounts of mucilaginous material (Fig. 3). Using EDX to assess elemental chemical composition of the spores (Fig. 4), only carbon and oxygen (approximately 50% total) was detected. Other elements present were not detected. There was no significant difference between the carbon results (p > 0.1). The A. niger 1988 spores recorded a weak amount of oxygen present (2.68%). Analysis of the substrata prepared “in house” (p(␥-MPS-COLMA)), (p(␥-MPS-MMA)) and PMMAsc was carried out using FTIR. Surfaces were composed of their expected chemistries (data not shown). Ra and contact angle measurements demonstrated a wide range of Ra values (358.4 nm–0.6 nm) and wettability measurements (108–42◦ ) (Table 1). There was no relationship between the Ra and the contact angle values. The surfaces were topographically diverse (Fig. 5): substrata with a Ra value above 32 nm for example PTFE (Fig. 5a) and glass (Fig. 5b) presented a range of irregular surface features whilst the remainder of surfaces showed very few topographical surface features (for example silicon Fig. 5c). The strength of attachment of A. niger 1957 to surfaces, was strongest to c-PMMA (24.2 nN) and weakest to (p(␥-MPS-MMA)) (9.5 nN) (Fig. 6a). For A. niger 1988 modified cantilevers, the strength of attachment was strongest to PMMAsc (43.1 nN) and

Aureobasidium pullulans

25 20 15 10 5 0 C

O

Na

Al

Si

K

Ti

Zn

Chemical element Fig. 4. EDX results demonstrating percentage weight of elements detected in fungal spores (n = 5).

weakest to PTFE (1.3 nN) (Fig. 6b). For Aureobasidium pullulans modified cantilevers the strongest attachment was to the glass surface (27.9 nN) and the weakest to the PTFE (0.4 nN) surface (Fig. 6c). The Aureobasidium pullulans spores attached with the widest range of forces to the surfaces (55.5 nN–0.36 nN), whilst A. niger 1957 attached with the smallest (24.28 nN–9.48 nN). All spores attached strongly to the glass and c-PMMA surfaces and weakly to the PTFE, (p(␥-MPS-co-MMA)) and (p(␥-MPS-co-LMA)) surfaces. For the A. niger 1957 and Aureobasidium pullulans the pattern of the strength of attachment to the surfaces was similar. The major difference in the A. niger 1988 results was that this spore demonstrated a strong attachment to the PMMAsc surface. When the force measurements were carried out using a blank cantilever on the substrata (Fig. 7),

Fig. 2. SEM images demonstrating different shapes and sizes of spores: (a) A. niger 1957, (b) A. niger 1988 and (c) Aureobasidium pullulans.

K.A. Whitehead et al. / Colloids and Surfaces B: Biointerfaces 82 (2011) 483–489

487

Fig. 5. Atomic force microscopy images demonstrating the differences in the surface topographies of the substrata: (a) PTFE, (b) glass and (c) silicon.

it was found that the strength of attachment did not always follow the trends observed with the spores. With the naked cantilever, force measurements using c-PMMA were comparable with those

Force (nN)

a

Force (nN)

b

Force (nN)

c

90 80 70 60 50 40 30 20 10 0

90 80 70 60 50 40 30 20 10 0

observed for the A. niger 1957 spore cantilever. Similar results were also observed for the naked cantilever or spore cantilevers with the (p(␥-MPS-co-LMA)). The naked cantilever also gave similar results on the PMMAsc surface to the A. niger 1988 spore cantilever. However, unlike the spore cantilever measurements, with the naked cantilever, the glass, PTFE and PMMAsc did not follow trends. There was no relationship found between the strength of attachment measurements of the spore probe cantilevers and the Ra values, but surface wettability influenced the results. 4. Discussion Cell adhesion is a complex process involving both physical and chemical events [40]. The complex structure and shapes of biological cells encompasses specific adhesions. When approaching a heterogeneous surface, spore:surface interactions result in a variety of long range, non-specific and short range specific interactions. Thus it is difficult to explain how the properties of the cell surface mediate attachment. There was little difference observed between the adhesion to hydrocarbons for the Aspergillus spp. whilst Aureobasidium pullulans spores adhered in weak numbers to the hydrocarbon phase. This may be due in part to the surface chemistry of the spores since Aureobasidium pullulans is known to excrete an exopolymeric substance, glucan which could be seen surrounding the spore using light microscopy. It is now believed that the MATH assay does not actually measure the surface hydrophobicity of the cells but an interplay of hydrophobic van der Waals, electrostatic interactions and of various short-range interactions [41]. Since the A. niger sp. had more affinity for the hexadecane, both species were arguably more easily wetted by non-polar media. This means that they have a more hydrocarbon like (hydrophobic) surface. The Aureobasidium pullulans was suspended in the aqueous phase and therefore has a more wettable (hydrophilic) surface [42] which may

90 80 70 60 50 40 30 20 10 0

Fig. 6. Strength of attachment of modified cantilever with (a) A. niger 1957, (b) A. niger 1988 and (c) Aureobasidium pullulans spores to surfaces. Error bars demonstrate standard error of the mean (n = 15).

Fig. 7. Strength of attachment of naked cantilever to surfaces. Error bars demonstrate standard error of the mean (n = 10).

488

K.A. Whitehead et al. / Colloids and Surfaces B: Biointerfaces 82 (2011) 483–489

be due to the presence of the mucilaginous material for example excreted exopolymeric substances (EPS). Thus this indirect method [43] revealed differences between the spore properties for the two genera investigated. SEM images of the spores revealed morphological differences but EDX was not sufficiently sensitive to determine a difference in chemical composition of the spores. Other methods such as X-ray photoelectron spectroscopy (XPS) or diffuse reflectance infra-red Fourier transform (DRIFTS) may be more useful in this instance. The surfaces presented a range of roughness and wettability measurements, but there was no relationship between the two parameters. This is not unexpected given the variety of chemistries and fabrication methods of the surfaces analysed. However, all surface topographies had Ra values below 200 nm. Fungal spores are large in size when compared to these surface features, and it is therefore questionable whether the small differences in Ra values would affect the strength of spore attachment. Work by Kearns and Barlocher, [44] on fungal spore attachment to leaf surfaces suggested that surface factors other than roughness influenced conidial attachment to leaf surfaces. The roughest surface was the commercially purchased PTFE whilst the smoothest was the silicon. The least wettable surfaces were the PTFE and (p(␥-MPSco-LMA)), whilst the most wettable surface was silicon, with the other surfaces ranging between, most of which ranged between water contact angles of 57–74◦ . Although the strength of attachment for the A. niger spores were similar despite their differing morphologies, the Aureobasidium pullulans spores attached to the test surfaces with a relatively wide range of force. The spherical A. niger 1957 spores demonstrated the least variability in results. A wider range was observed for the spikey A. niger 1988 spore, thus although the spores are the same genus the morphology (i.e. spikes) influenced the strength of attachment. This is presumably due to the increased heterogeneity of the surface conveyed by the surface projections. The range shown by Aureobasidium pullulans could be due to either the presence of EPS, or to the large and/or irregular surface shape of the spore. Both factors will contribute to spore surface/shape heterogeneity at a level beyond the resolution available to this study. All the spore types attached strongly to all the wettable surfaces. The lack of attachment to the PTFE and (p(␥-MPS-co-LMA)) will be due to the strong surface water repulsion of these materials. All the modified spore cantilevers attached weakly to PTFE and (p(␥-MPS-co-LMA)) (non-wettable) and (p(␥-MPS-co-MMA)) surfaces. Although these results were expected for the PTFE and (p(␥-MPS-co-LMA)) since they were the most non-wettable surface, one might anticipate similar interactions with the c-PMMA, (p(␥-MPS-MMA))and the PMMAsc due to similar chemistry, wettability and Ra values, [the Ra value on the (p(␥-MPS-MMA)) surface is one log higher than the others due to a waviness of form rather than increased surface topography]. However, the fabrication method may have an effect. The c-PMMA was bulk polymerised between glass sheets, whereas the PMMAsc was prepared using a solution of MMA in a solvent, which was spin coated onto a silicon wafer with the solvent evaporated off. The structure and mobility of the chemical moieties of the surface will be dependent on the concentration and viscosity of the solution, and thus the surface structure of this material may be more dynamic than that of the c-PMMA. The dynamic nature of the surface may be the reason for the unusual results seen for the three spores. The (p(␥-MPS-MMA)) was produced by the adsorption of the silane ␥-MPS onto a silicon wafer followed by copolymerisation with MMA. This is likely to produce branched structures which will have a strong molecular mobility. When comparing the results of the spore cantilevers with the results of the naked cantilever it may be suggested that specific interactions with the surfaces dominate since each spore cantilever type tested gave a different range of strength of attachment mea-

surements and individual binding profiles. It may be that specific binding interactions with the complex surface structures and binding motifs of the fungal spores influence the strength of spore attachment to the substrata. However, this needs to be further investigated. Spore shape (A. niger 1988) rather than wettability affected the trend in results. A. niger 1988 demonstrated a strong strength of attachment to the PMMAsc surface which was not observed with the other spore types. This may be due to the surface projections of these spores interacting favourably with the PMMAsc surface. There was no trend observed between the surface roughness and strength of spore attachment to the surfaces, but this will have occurred since unlike most attachment assays the spore is only momentarily brought into vertical contact with the surface, and thus surface roughness does not have a significant effect on the results. The force distance curves generated during this work demonstrated evidence of direct adhesion of the spore cantilever to the surface. This may be explained since the spore coats are extremely hard and are only in contact with the surface for a fraction of a second, thus evidence of polymer stretching was not observed. A number of studies have assessed the role of physicochemical surface properties in the attachment of bacterial spores [45–47]. Using Geobacillus sp. spore adhesion to a range of surfaces Seale et al. [19] found no simple relationship between individual physicochemical interactions and spore adherence; they also determined that surface modifications which limit the attachment of one strain may not be effective for all strains of the same species and control regimes need to be devised with reference to the characteristic of the particular strains of concern [19]. 5. Conclusions In conclusion, spores from two strains of A. niger and one strain of Aureobasidium pullulans attached strongly to wettable c-PMMA and glass surfaces, and weakly to the (p(␥-MPS-coMMA)) and non-wettable PTFE and (p(␥-MPS-co-LMA)) surfaces. All three strains interacted similarly with surfaces. The dynamic nature of the (p(␥-MPS-co-MMA)) surface may also have affected results. No relationship was demonstrated between the strength of attachment of the spores to the surfaces and the roughness measurements of the surfaces. However, surface wettability did influence the strength of spore attachment to surfaces but not in an easily interpretable manner. This work has demonstrated that in strength of attachment studies surface wettability, but not roughness influenced spore attachment to the surfaces. The shape of the spore (projections) also influenced results. The findings from this work will help elucidate the interactions between fungal spores and substrata and enhance the selection of surfaces for uses in environments were biofouling/biodeterioration of substrata is an important consideration. Acknowledgements This work was supported by Ciba Speciality Chemicals Corporation. The authors would like to thank Todd Elder and Tom Xiuang (Ciba Speciality Chemicals Corporation, New York) for their contribution in the production of the spin coated surfaces used in this study. References [1] [2] [3] [4]

M.J. Dempsey, Mar. Biol. 61 (1981) 305. K.C. Marshall, R. Stout, R. Mitchell, Can. J. Microbiol. 17 (1971) 1413. J.W. Gillatt, Jocca-Surf. Coat. Int. 74 (1991) 197. T. Warscheid, J. Braams, Int. Biodeter. Biodeg. 46 (2000) 343.

K.A. Whitehead et al. / Colloids and Surfaces B: Biointerfaces 82 (2011) 483–489 [5] O.M. Zacheus, M.J. Lehtola, L.K. Korhonen, P.J. Martikainen, Water Res. 35 (2001) 1757. [6] G. Magaudda, J. Cultural Heritage 5 (2004) 113. [7] R. Purwar, M. Joshi, AATCC Rev. 4 (2004) 22. [8] C.B. Greenberg, C. Steffek, Thin Solid Films 484 (2005) 324. [9] S. Scheerer, O. Ortega-Morales, C. Gaylarde, Adv. Appl. Microbiol. 66 (2009) 97. [10] V. Jurado, S. Sanchez-Moral, C. Saiz-Jimenez, Int. Biodeter. Biodeg. 62 (2008) 325. [11] S.R. Barratt, A.R. Ennos, M. Greenhalgh, G.D. Robson, P.S. Handley, J. Appl. Microbiol. 95 (2003) 78. [12] F. Cappitelli, C. Sorlini, Appl. Environ. Microbiol. 74 (2008) 564. [13] L. Cosgrove, P.L. McGeechan, G.D. Robson, P.S. Handley, Appl. Environ. Microbiol. 73 (2007) 5817. [14] H. Ma, C.J. Winslow, B.E. Logan, Colloids Surfaces B: Biointerfaces 62 (2008) 232. [15] W.R. Bowen, R.W. Lovitt, C.J. Wright, J. Colloid Interface Sci. 228 (2000) 428. [16] J. Clement, S. Martin, R. Porter, T. Butt, A. Beckett, Mycol. Res. 97 (1993) 585. [17] J. Hamer, R. Howard, F. Chumley, B. Valent, Science 239 (1988) 288. [18] P. Volz, Microbios 91 (1997) 145. [19] R.B. Seale, S.H. Flint, A.J. McQuillan, P.J. Bremer, Appl. Environ. Microbiol. 74 (2008) 731. [20] W. Cooke, Mycopathol. Mycol. Appl. 12 (1959) 1. [21] D. Eveleigh, Ann. Appl. Biol. 49 (1961) 403. [22] N. Hamilton, in: T. Oxley, S. Barry (Eds.), Biodeterioration 5, John Wiley and Sons, Chichester UK, 1983, p. 663. [23] J.M. Pouliot, I. Walton, M.N. Parkhouse, L.I. Abu-Lail, T.A. Camesano, Biomacromolecules 6 (2005) 1122. [24] J. Andrews, R. Harris, R. Spear, W. Gee, E. Nordheim, Can. J. Microbiol. 40 (1994) 6. [25] S.L. Bardage, J. Bjurman, Can. J. Microbiol. 44 (1998) 954. [26] R.D. Boyd, J. Verran, M.V. Jones, M. Bhakoo, Langmuir 18 (2002) 2343.

489

[27] G. Midelet, A. Kobilinsky, B. Carpentier, Appl. Environ. Microbiol. 72 (2006) 2313. [28] S.G. Parkar, S.H. Flint, J.S. Palmer, J.D. Brooks, J. Appl. Microbiol. 90 (2001) 901. [29] K.A. Whitehead, D. Rogers, J. Colligon, C. Wright, J. Verran, Colloids Surfaces B: Biointerfaces 51 (2006) 44–53. [30] J. Verran, R.L. Taylor, G.C. Lees, J. Mater. Sci.: Mater. Med. 7 (1996) 597. [31] J. Webb, H. Van der Mei, M. Nixon, I. Eastwood, M. Greenhalgh, S. Read, G. Robson, P. Handley, Appl. Environ. Microbiol. 65 (1999) 3575. [32] M. Rosenberg, S. Kjelleberg, Adv. Microbial Ecol. 9 (1986) 353. [33] J. Verran, R.D. Boyd, K. Hall, R.H. West, J. Food Prot. 64 (2001) 1377. [34] J.F. Frank, R.A.N. Chmielewski, J. Food Prot. 60 (1997) 43. [35] D.R. Korber, A. Choi, G.M. Wolfaardt, S.C. Ingham, D.E. Caldwell, Appl. Environ. Microbiol. 63 (1997) 3352. [36] J. Verran, K.A. Whitehead, ICHEME Food Bioprod. Process. 84 (2006) 260. [37] W.R. Bowen, R.W. Lovitt, C.J. Wright, Colloids Surfaces A: Physicochem. Eng. Aspects 173 (2000) 205. [38] W. Ducker, T. Senden, R. Pashley, Nature 353 (1991) 239. [39] M. Rosenberg, R.J. Doyle, in: R.J. Doyle, M. Rosenberg (Eds.), Microbial Cell Surface Hydrophobicity, American Society for Microbiology, Washington, D.C., USA, 1990, p. 1. [40] M. Berglin, A. Olsson, H. Elwing, Macromol. Biosci. 8 (2008) 410. [41] H.C. van der Mei, B. van de Belt-Gritter, H.J. Busscher, Colloids Surfaces B: Biointerfaces 5 (1999) 117. [42] R. Bos, H.C. van der Mei, H.J. Busscher, FEMS Microbiol. Rev. (1999) 179. [43] G. Geertsema-Doornbusch, H. van der Mei, H. Busscher, J. Microbiol. Methods 18 (1993) 61. [44] S.G. Kearns, F. Barlocher, Fungal Ecol. 1 (2008) 13. [45] C. Faille, C. Jullien, F. Fontaine, M. Bellon-Fontine, C. Slomianny, T. Benezech, Can. J. Microbiol. 48 (2002) 728. [46] U. Husmark, U. Ronner, Biofouling 5 (1992) 335. [47] U. Husmark, U. Ronner, Biofouling 7 (1993) 57.