New Developments in Coatings Technology - American Chemical

Chapter 7

Laboratory Methods to Access the Antifouling and Foul-Release Properties of Novel, Non-Biocidal Marine Coatings 1

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Downloaded by UNIV OF ARIZONA on January 19, 2013 | http://pubs.acs.org Publication Date: August 30, 2007 | doi: 10.1021/bk-2007-0962.ch007

M . E . Callow , J . A . Callow , and D. E . Wendt

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School of Biosciences, University of Birmingham, Birmingham B15 2TT, United Kingdom Biological Sciences Department and Center for Coastal Marine Sciences, California Polytechnic State University, San Luis Obispo, C A 93407

In this article the advantages and disadvantages o f laboratory-based methods to assess the performance o f antifouling and foul-release coatings for application in the marine environment are discussed. Properties o f the major groups o f macrofouling organisms (algae and invertebrates) are briefly considered before the various assay methods are outlined. The nature o f results obtained is illustrated by reference to sample data.

© 2007 American Chemical Society

In New Developments in Coatings Technology; Zarras, P., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 2007.

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Introduction Biofouling is the accumulation o f microorganisms, algae and animals on man-made surfaces. The marine environment is unique because o f the diversity o f organisms that cause biofouling. Within minutes o f immersing a clean surface in water it becomes 'conditioned' by adsorption o f a macromolecular film. Bacteria colonise within hours, as may unicellular algae, cyanobacteria (blue-green algae) protozoa and fungi. These early small colonisers form a biofilm, which is an assemblage o f attached cells often referred to as 'microfouling' or ' s l i m e ' . Macrofouling consists o f multicellular algae (i. e. the 'seaweeds'), soft-bodied invertebrates such as sponges and tunicates ('soft' macrofouling), and calcified invertebrates such as barnacles and tubeworms ('hard' macrofouling). Although a microbial biofilm may moderate the settlement o f spores and larvae o f more complex organisms, it is not a pre-requisite (7). O n ships, most interest has focused on fouling by barnacles but other important invertebrate foulers are tubeworms, bryozoans, hydrozoans and sponges. Ulva (syn. Enteromorpha) is the most important macroalga because it is ubiquitously distributed and can withstand fluctuating environmental conditions. Microfouling shines are dominated by diatoms which are o f interest because they are difficult to remove from non-biocidal, silicone-based foulrelease coatings (2). A l l fouling organisms secrete 'sticky' materials (adhesive proteins, glycoproteins and polysaccharides) to attach to substrata (3). Larvae o f invertebrates and spores o f algae need to quickly locate and attach to a surface in order to complete their life history. Non-biocidal coatings can be manipulated to deter settlement through for example, impressed microtopographic features (4). Controlling fouling without the use o f biocides is essentially a problem o f deterring settlement and/or controlling adhesion. T o assess the potential o f foul-release coatings, which release fouling under robust hydrodynamic conditions, necessitates the use o f methods that measure the adhesion strength o f attached organisms.


93 In this paper, we review the main laboratory-scale methods used to evaluate non-biocidal coatings for antifouling and fouling release properties. The key test organisms are the cypris larvae and young adult (juvenile) stages of barnacles, and the spores and young plants (sporelings) of the green alga Ulva.

Why is there a need for laboratory-scale evaluations of


coatings? This question is asked frequently. O n the positive side, laboratory testing allows a rapid assessment o f a coating or surface under controlled and reproducible conditions, thus the challenge is not site-specific or seasonal in character. Laboratory-scale evaluation is relatively low cost, both in terms o f facilities, and the amount o f test material required, an important factor where novel polymer systems are being investigated at an experimental scale. Furthermore, it allows the different stages o f settlement, attachment and fouling release to be studied in detail. This is especially important, as the key to antifouling (cf. weak adhesion and fouling release) necessitates control o f the settling stages (spores or larvae), which are not considered in field tests employing panels hung from rafts. Issues relating to film stability and bonding to the substrate also become apparent at an early stage the consequences o f which may render expensive and time-consuming field trials worthless. In terms o f coatings development, laboratory assays allow model systems to be evaluated that provide valuable data to synthetic chemists in terms o f informing coatings development. Information from a failure can be just as valuable as from a success i f it is possible to determine why a coating failed. O n the negative side, it can be argued that laboratory assays do not reflect the real world, where changing environmental conditions, season, competition and prédation strongly influence the biofouling community structure as well as more subtle factors such as the adhesion strength o f a particular organism. Assays with


94 a single species may miss synergistic effects present in more com plex communities


Key test organisms. 1: the green seaweed Viva Mature, reproductive plants release vast numbers o f motile, unicellular, pear-shaped spores (technically 'zoospores'), 5-8 μηι in length (Plate la). Swimming spores attach rapidly to a surface once the spore has 'detected" that it is suitable for settlement. Secretion o f adhesive (Plate lb) results in firm attachment to the substratum. The adhesive cures rapidly by some form o f cross-linking (5) which is manifest as increased adhesion strength with time after settlement (6). The spores germinate and grow into young plants (sporelings, Plate l e ) , a few hundred microns in length, within a few days.

Key test organisms 2: Barnacles Balanus amphitrite is the most common species o f barnacle utilized in laboratory assays. It is easy to culture, fast-growing, and cosmopolitan in geographic distribution. Mature adults release thousands o f free-swimming larvae ('nauplii'). The larvae actively feed and undergo a total o f 6 instar stages. The final larval stage is the non-feeding cypris larva (Plate Id). The cypris larva is approximately 500μηι in length and is specialized for attachment and metamorphosis. It has a pair o f sensory antennae that are used for determining physical and chemical properties o f the substratum. The cypris larva explores the substratum by ' w a l k i n g ' on its antennae. When the cypris larva identifies a suitable substratum, it attaches by secreting an adhesive from cement glands located at the base o f the antennae. After attachment the larva undergoes a complex metamorphosis into a juvenile barnacle and thence into an adult (Plate le).


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Test sample format The standard format for the assays described below is coated glass microscope slides. Acid-washed slides are usually included in assays as standards. Smooth coatings of uniform thickness, without sagging or 'picture framing' and well-bonded to the glass are essential if hydrodynamic studies in a flow channel apparatus are to be performed. Some coatings that contain toxic curing agents e.g. dibutyltin compounds, need to be leached before the assays are set up. This is usually accomplished in a tank of recirculating deionised water fitted with a carbon filter. Leachates are collected and monitored by bioassay (7) until no toxicity is measured. The slides are transferred to artificial seawater (ASW) for at least 1 hour before an assay is set up.

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' Antifouling Settlement Assays Assays with Algae Ulva plants, collected from the seashore, release zoospores on immersion in seawater. The spore concentration is adjusted to 1-2 xlO / ml (using absorbance at 660 nm). Six replicates of each coating are required if spore settlement and strength of attachment assays are to be performed; three for settlement and three for adhesion strength. For sporehing growth and strength of attachment assays, a further six replicates are needed. Slides are placed in individual compartments of Quadriperm dishes (Fischer) to which 10 ml zoospore suspension are added. The dishes are incubated for 1 h in darkness at room temperature (c. 20°C). Slides are gently washed in ASW to remove unattached spores and fixed in 2.5% glutaraldehyde in seawater (vlv). Slides are air dried after washing in distilled water. Spore numbers are usually counted (30 counts per slide) at 1 mm intervals across the middle of the slide using a Zeiss Kontron 3000 image analysis system attached to a Zeiss epifluor6



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Plate 1. a) Zoospores of the green alga U l v a showing the 4 locomotory flagella (false-colour scanning electron micrograph image), b) false-colour image of a settled spore of Ulva as seen by Environmental Scanning Electron Microscopy. The original spore body is now surrounded by an annulus of secreted, hydrogel-like adhesive. The flagella are absorbed during settlement. (Copyright 2005from Walker et al, Journal of Adhesion 81,1101-1118; reproduced by permission of Taylor & Francis Group, LLC, http:llwww.taylorandfrancis.com). c) young plant ('sporeling') of U l v a which develops from germinated settled spores, d) The cypris larva of the barnacle Balanus amphitrite showing two anterior antennules (arrowed) and numerous posterior thoracic appendages usedfor swimming. Also apparent are multiple lipid droplets, the larval energy stores, e) top view of a young adult barnacle ofB. amphitrite, which metamorphosedfrom the settled cypris larva, f) bottom view of an adult barnacle attached to a clear silicone coating showing the basal plate. (See page 9 of color inserts.)


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escence microscope via a video camera. Spores are visualised by autofluorescence of chlorophyll.


Assays with barnacles

Antifouling settlement assays using barnacles (or larvae of the other invertebrates) are conducted using a "drop assay" or in certain situations a settlement preference experimental design. Which assay is utilized depends on the physical nature of the surface being tested. For example, a drop will not form on extremely hydrophobic or hydrophilic surfaces and thus the coatings need to be continuously submerged for testing. It is recommended that at least 5 replicate experimental surfaces and an additional 5 replicate control surfaces, uncoated glass slides, are used in antifouling assays. For the drop assay, a 0.4ml aliquot of seawater containing 2050 cyprids is placed on the surface of each replicate slide. The slides are placed in covered Petri dishes with moistened paper towels. The Petri dishes are placed in a constant temperature (25°C) and light (12 hour light-dark cycle) incubator. The assay lasts until 50% of the cyprids on the control surfaces have attached, or until 72 hours have elapsed. At the conclusion of the assay, the number of swimming larvae and the number of attached barnacles is enumerated for each replicate surface. Percent attachment is calculated by comparing the number of settled barnacles to total number of cyprids exposed to the surface. The settlement preference assay differs from the drop assay in that larvae are presented with a choice: the experimental coating or the polystyrene surface of the Petri dish in which the coating is submerged. The basic protocol consists of adding 50-100 larvae in 40 ml of seawater to a Petri dish (100 χ 15mm) containing an experimental or control slide. The duration of the experiment is determined as described above. At the conclusion of the assay, the number of swimming cypris larvae and the number of attached barnacles on the slide and Petri dish are enumerated. The percentage of individuals that settled on the slide and on the Petri


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dish are calculated. From these data a settlement preference ratio is calculated: the number of settled barnacles per available square millimeter of the experimental coating divided by the number of settled barnacles per available square millimeter of the Petri dish. A settlement preference ratio of 1 indicates no preference for either surface, whereas a settlement preference ratio less than one indicates a coating that may deter settlement.


Assays to evaluate foul-release potential

Assays with algae

Slides settled with Ulva zoospores are exposed to turbulent flow in a specially designed flow channel or exposed to the compressive forces of a water jet. The design and operating principles of the two apparatuses are described below. Cell density on three replicate slides before and after exposure is determined as described above. Percentage removal data are calculated from the mean number of spores remaining attached to the surface after exposure to turbulent flow (flow channel) or surface pressure (water jet) compared with the mean number before the slides were subjected to flow/water pressure (e.g. 8,9,10). Foul-release potential can also be assessed on Ulva sporelings (young plantlets formed from germinated settled zoospores after a few days growth). Zoospores are settled on 6 replicates of each test surface as described above. The washed slides are replaced in the Quadriperm dishes and 10 ml of growth medium added to each dish compartment (8). The dishes are incubated at 18 °C with a 16 h:8 h, light: dark cycle, the medium being refreshed every 2 days. After 78 days, bioinass on the surface of the slides is quantified before and exposure to flow in the flow channel. Biomass is recorded either directly in a TECAN plate reader using the autofluorescence of chlorophyll, or the sporelings are harvested by scraping with a razor blade. In the latter case, biomass is determined as extracted


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chlorophyll. A foul-release silicone elastomer, e.g. Silastic T2 (Dow Corning) is normally included as a positive standard whilst glass is included as a negative standard.


Assays with barnacles

Laboratory evaluation of foul-release properties of coatings using barnacles is done using an automated test stand that applies a shear force to animals attached to coatings (77). Barnacles with a basal plate diameter >3-5 mm are used as basal plate breakage occurs more frequently with smaller animals (72). Prior to dislodgement tests, the basal plates of barnacles (Plate If) are photographed with a digital camera mounted on a dissecting microscope. Digital analysis (NIH. ImageJ) is used to calculate the area of the basal plate. The coated slide is then clamped in the force-gauge testing apparatus. Data can be streamed continuously to a computer which allows creation of a force vs. time curve for analysis of fracture processes. Peak force is also recorded and is used to calculate critical removal stress (force per unit area) for each barnacle. In the event that the barnacle basal plate fractures during testing, the force at which it broke is recorded, and the percentage of area of the basal plate remaining attached is determined via digital image analysis. In addition to direct comparisons of critical removal stress, the linear relationship between basal plate area and percentage of basal plate remaining on a coating is a measure of the efficacy of foul-release surfaces (72).

Apparatus

The fully turbulent flow channel and water jet apparatus

The operating principle (75) is that of a high aspect ratio, turbulent channel flow since boundary layers around ships are


101 turbulent in character. The fully-developed channel flow allows determination o f accurate wall shear stress from a simple pressure gradient measurement. The flow channel holds 6 microscope slides with or without coatings. Turbulent flow is created in a 60 c m long low aspect ratio section o f channel preceding the slides. Flows o f A S W up to 4.9 m s" generate wall shear stresses up to 56 Pa. The relationship between three models and the results from laboratory scale tests to the self-cleaning o f a ship coated with a foul-release surface are discussed in (14). The water jet developed by Swain & Schultz (75), designed for assessment o f fouled panels in the field, uses perpendicular rather than parallel flow and delivers a greater force/unit area than the flow channel. Modifications to the original field apparatus have produced an automated, standardized operation by computer-driven stepper motors that allow the jet nozzle to be raster-scanned across a batch o f slides at a controlled rate, in a variety o f reproducible patterns (6). The instrument is typically operated at a speed o f 10 mm s" for 10 swathes, at the end o f which an area o f 500 m m in the mid-region o f each slide has been exposed to the jet o f water. The water supply is housed in a pressure resistant tank and pressurised using a compressed air supply from a conventional S C U B A tank. The relationship between regulator setting and impact pressure exerted at the surface is described in (75).


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Force Gauge Test Stand Removal force testing with barnacles is done using an automated test stand ( I M A D A model number S V - 5 ) and a digital force gauge ( I M A D A model number A X T - 7 0 o z R ) (77). The capacity o f the force gauge is up to 19.5 Ν with a sensitivity o f 0.01 N . The force gauge is mounted on a motorized arm that allows for a constant rate o f application at 60-75 microns/second. The reproducibility o f angle and rate o f application are critical to reducing variability between measurements. The test stand has a custom-made acrylic chamber in which test coatings are clamped.


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The chamber ensures that the barnacles are continuously immersed in seawater during dislodgment tests. Data from the force gauge can be streamed using an RS-232 digital interface at 20 recordings/second, which allows determination of force vs. time curves as well as maximum force during dislodgement.


Examples of Lab-scale Evaluation The application of these methods to the evaluation of actual test samples can be illustrated by reference to recent published data. Gudipati et al. (S) used Ulva to explore antifouling and foul-release properties of a series of cross-linked hyperbranched fluoropolymers (HBFP) and poly(ethylene glycol) (PEG) amphiphilic networks. Settlement of Ulva zoospores was consistently and markedly lower on all the cross-linked HBFP-PEG network formulations compared to glass controls (Figure 1). Removal of spores from the HBFPPEG45 coating in the flow channel was significantly different from the rest (Figure 2). Greatest removal of sporelings was also obtained from the HBFP-PEG45 surface with a mean detachment of over 90% (Figure 3). This was approximately twice of that from the T2 Silastic PDMS standard which suggests that this nanostructured material has promising properties in the practical control of soft fouling. The best illustration of the settlement preference assay using barnacles comes from unpublished data on proprietary coatings. The protocol can clearly resolve differences in settlement preference between coatings as is illustrated by vastly different ratios between surfaces (Figure 4). Tang et al. (9) used B. amphitrite to evaluate the foul-deterrent and foul-release properties of experimental hydrophobic and hydrophilic xerogel films. Settlement rates of cypris larvae on xerogels containing C3-TMOS, C8-TEOS, and enTMOS were lower than that of larvae on glass and polystyrene controls, but not significantly so (Figure 5). The 50/50 C8-TMOS/TMOS was significantly lower than glass controls. In the same study critical removal stress was evaluated for 50/50 C8-TEOS/TMOS xerogel coatings with different sol-



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Figure 1. Settlement of Ulva spores upon cross-linked HBFP-PEG network coatings. (Reproducedfrom reference 8. Copyright 2005 American Chemical Society.)

Figure 2. Removal of attached Ulva spores from HBFP-PEG networks in the flow apparatus (55 Pa wall shear stress). Removal was greatest from the coating with 45 wt % PEG, indicating its foul-release properties. (Reproduced from reference 8. Copyright 2005 American Chemical Society.)



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Figure 3. Percentage removal of Ulva sporelings in the flow apparatus. The crosslinked HBFP-PEG coating with 45 wt% PEG was more effective in fouling release than the standard PDMS coating. (Reproducedfrom reference 8. Copyright 2005 American Chemical Society.)

Figure 4. Settlement preference of cypris larvae ofB. amphitrite exposed to proprietary coatings. A settlement preference ratio of indicates no preference for either surface, whereas a settlement preference ratio

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