Polymer International Polym Int 57:1042–1051 (2008)

Surface changesin polyhydroxyalkanoate films duringbiodegradation and biofoulingCatherine A Woolnough,1 Tim Charlton,2 Lachlan H Yee,2 Maria Sarris3 andL John R Foster1∗1Bio/Polymer Research Group and Centre for Advanced Macromolecular Design, School of Biotechnology & Biomolecular Sciences,University of New South Wales, Sydney, NSW 2052, Australia2Centre for Marine Bio-Innovation, University of New South Wales, Sydney, NSW 2052, Australia3Histology and Microscopy Unit, School of Medical Sciences, University of New South Wales, Sydney, NSW 2052, Australia

Abstract

BACKGROUND: Despite the recognition that microbial biofilms play a role in environmental degradation ofbioplastics, few studies investigate the relationship between bioplastic biodegradation and microbial colonisation.We have developed protocols based on a combination of confocal laser scanning microscopy and contact anglegoniometry to qualitatively and quantitatively map surface changes due to biofilm formation and biopolymerdegradation of solvent cast poly(3-hydroxyalkanoate) films in an accelerated in vitro biodegradation system.

RESULTS: A significant regression relationship between biofilm formation and polymer biodegradation(R2 = 0.96) was primarily conducted by cells loosely attached to the film surfaces (R2 = 0.95), rather than thestrongly attached biofilm (R2 = 0.78). During biodegradation the surface rugosity of poly(3-hydroxybutyrate) andpoly[(3-hydroxybutyrate)-co-(3-hydroxyvalerate)] increased by factors of 1.5 and 1.76, respectively. In contrast,poly(3-hydroxyoctanoate) films showed little microbial attachment, negligible weight loss and insignificant changesin surface rugosity.

CONCLUSION: A statistically significant link is established between polymer weight loss and biofilm formation.Our results suggest that this degradation is primarily conducted by cells loosely attached to the polymer ratherthan those strongly attached. Biofilm formation and its type are dependent upon numerous factors; the flatundifferentiated biofilms observed in this study produce a gradual increase in surface rugosity, observed as anincrease in waviness. 2008 Society of Chemical Industry

Keywords: polyhydroxyalkanoates; biodegradation; biofilms; surface roughness; confocal laser scanningmicroscopy

INTRODUCTIONPolyhydroxyalkanoates (PHAs) are a diverse groupof biopolyesters, accumulated intracellularly by awide range of microorganisms. PHAs function pri-marily as carbon and energy reserves.1,2 Poly(3-hydroxybutyrate) (PHB) is the most common mem-ber of the PHA family and has commercial poten-tial as a biodegradable replacement for persis-tent conventional plastics. However, due to itsphysicochemical properties, melt pressed or sol-vent cast films of PHB are brittle materials.3

Copolymerisation of 3-hydroxybutyrate (HB) withunits of 3-hydroxyvalerate (HV) to yield poly[(3-hydroxybutyrate)-co-(3-hydroxyvalerate)] [P(HB-co-HV)] modifies the physicochemical properties of suchfilms making them more flexible, leading to thiscopolymer’s application as an environmentally friendly

bioplastic.3,4 In contrast, poly(3-hydroxyoctanoate)(PHO) is a PHA possessing medium length alkylgroups in the side chain (mcl-PHA) and is usu-ally a terpolymer of C6, C8 and C10 monomericunits. Compared to the short chain length PHAs (scl-PHA), PHB and P(HB-co-HV), PHO has a lowercrystallinity, a lower melting temperature and greaterflexibility.5

The environmental degradation of PHB and P(HB-co-HV) has been reported in a variety of naturalenvironments, while environmental degradation ofPHO is limited to the report of Tan and coworkers.6,7

Biodegradability of PHB and P(HB-co-HV) is influ-enced by the valerate content, the degree of crys-tallinity, the microbial population and other abioticparameters. Generally, a higher degree of crystallinityleads to a lower rate of biodegradation with preferential

∗ Correspondence to: L John R Foster, Bio/Polymer Research Group, School of Biotechnology & Biomolecular Sciences, University of New South Wales,Sydney, NSW 2052, AustraliaE-mail: J.Foster@unsw.edu.au(Received 6 February 2008; revised version received 9 April 2008; accepted 14 May 2008)Published online 14 July 2008; DOI: 10.1002/pi.2444

2008 Society of Chemical Industry. Polym Int 0959–8103/2008/$30.00

Biodegradation and biofouling of PHA films

degradation of the amorphous phase.8 Thus, P(HB-co-HV) was found to degrade faster than PHB infresh water and marine environments as well as insoil.2,9 There are limited numbers of studies compar-ing biodegradation rates of PHB, P(HB-co-HV) andPHO. While PHO is more amorphous than its scl-PHAcounterparts, its rate of biodegradation is approxi-mately 22 times slower than that of PHB in mangrovesoil.7 Although microbial ecology is recognised as play-ing an important role in environmental degradation ofbioplastics, biodegradation studies tend to focus onthe physicochemical and mechanical properties of thematerial. However, there is an increasing recogni-tion of the environmental and medical importance ofbiofilms; this has led to studies investigating the for-mation of biofilms on polymers such as polyethyleneand rubber, as well as their influence on the possiblebiodegradation of such materials.10,11 Related studieshave investigated the addition of antimicrobial com-pounds to biodegradable polymers to prevent biofilmformation in medical settings.12

Various surface characteristics influence materialcolonisation by microorganisms and the formation ofbiofilms. These include surface roughness, hydropho-bicity, charge, microbial population and, with particu-lar reference to this study, whether or not the materialcan be degraded by the microorganisms in a biofilm.Rough surfaces may be easier to colonise, as cellsin crevices are protected from adverse sheer forces.13

Furthermore, rough surfaces provide a greater surfacearea to bulk ratio, facilitating increased biofilm for-mation. Pasmore et al.14 have reported that increasedroughness, hydrophobicity and charge leads to greaterbiofilm coverage of polyethylene, Teflon and cel-lulose acetate. In contrast, Eginton et al.15 showedthat bacterial colonisation of glass, poly(vinyl chlo-ride), polystyrene, stainless steel and Formica was notrelated to hydrophobicity or roughness, although thestrength of attachment was dependent upon surfacehydrophobicity. The study of the interaction of biofilmformation, surface modification and biodegradation ofpolymeric materials is still in its infancy and requiresfurther study.

Previous surface studies of PHB and P(HB-co-HV)during their degradation have been limited to qualita-tive comparisons of undegraded and partially degradedmaterials using scanning electron microscopy whichrevealed apparent increases in roughness duringdegradation, based on the visual extent of surface‘pitting’.16,17 While PHB and P(HB-co-HV) biopoly-mers are known to be environmentally degradable

and susceptible to various extracellular depolymerises,there are virtually no studies investigating biofilmformation on these biomaterials. Biofilm formationon these biopolymers is of particular interest giventheir potential as biodegradable replacements for con-ventional persistent plastics. Hydrolytic degradationof PHB is a surface-driven phenomenon, with anumber of studies demonstrating that degradationis partly dependent on the surface area to bulkproperties.16–18 It may be possible, through modi-fication of the surface properties such as roughnessand hydrophobicity, to control biofilm formation onthese bioplastics and consequently their degradationrates.

Biofilms form on most surfaces exposed to environ-ments possessing biological entities; in the process,microbial colonisation can be rapidly compounded bythe attachment of fungi, algae and higher organisms.19

Consequently, for standardisation purposes biofilmstudies tend to focus on the initial microbial colonisa-tion using in vitro experiments.19,20

In this study, an accelerated in vitro biodegrada-tion model containing an enriched, undefined, mixedculture of bacteria from soil was developed. Fur-thermore we report on the application of confocallaser scanning microscopy (CLSM) and contact anglegoniometry (CAG) combined with fluorescent stain-ing to scl-PHA and mcl-PHA solvent cast films tomonitor biofouling and surface rugosity. The paperdemonstrates the application of these techniques toqualitatively and quantitatively monitor changes insurface parameters including biofilm colonisation dur-ing degradation.

MATERIALS AND METHODSMaterialsPHB of microbial origin and its copolymer with8 mol% HV units, P(HB-co-8HV), were purchasedfrom Sigma Aldrich (Sydney, Australia; product num-bers 363502, 403113 and batch numbers 27331CS,12703MD, respectively) and used as supplied. PHOwas produced using Pseudomonas oleovorans (ATCC29347) according to Foster et al.21 PHA films werecharacterised using gas chromatography (GC), NMRspectroscopy, gel permeation chromatography (GPC)and X-ray crystallography as described previously;Table 1 summarises the composition, molecular massand crystallinity of the samples used in this study.21 Allother chemicals were obtained from APS Chemicals(Seven Hills, Australia) and used as supplied.

Table 1. Composition, molecular properties and crystallinity of solvent cast PHA films

Composition (mol%)

PHA C4 C5 C6 C8 C10 Mw (×103 g mol−1) Mn (×103 g mol−1) D Crystallinity (%)

PHB 100 600 240 2.5 88P(HB-co-8HV) 92 8 252 103 2.4 76PHO 6 92 2 235 100 2.4 34

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Polymer film fabricationPHB, P(HB-co-8HV) and PHO were dissolved inchloroform and fabricated into films by casting intoclean, dry, glass Petri dishes in a fume hood. Two glasscovers were placed over the dishes and the chloroformevaporated slowly over 4 days. The films were driedfor 48 h in a vacuum desiccator and removed from thedishes and allowed to stand for a further 24 h untiltheir weights had stabilised in air. The films were agedfor a further three weeks to enable their crystallinityto reach equilibrium. Films were subsequently cutinto 8 × 8 mm pieces approximately 200 µm thickand weighing around 20 ± 3 mg (Cahn C-33, Orion,USA, microgram balance, accuracy of ±10 µg). Forstandardisation purposes all microscopic imaging wasperformed on film sides that were cast in contact withthe clean glass.

Preparation of mixed microbial cultureBacteria were extracted from fertile garden soilcollected at the Ecoliving Centre, University of NewSouth Wales, Sydney, Australia (pH = 6.5, watercontent 16% and temperature 15 ◦C) and enrichedusing a protocol reported by Yu.22 Briefly, this wascarried out by soaking 0.5 kg of soil per litre of tapwater for 1 h then filtering the suspension throughNo. 1 Whatman filter paper. An amount of 40 mL ofthe filtrate was added to 60 mL of media containing2 g glucose, 2 g yeast extract and 2 g peptone per litreand incubated at 25 ◦C for 24 h. Microorganisms inthese cultures were then collected via centrifugation(6000 × g, 10 min) and added to minimal mediawith 700 mg L−1 of PHB films, 700 mg L−1 ofP(HB-co-8HV) films and 700 mg L−1 of PHO films asthe carbon sources. The minimal media consistedof K2HPO4 (3.76 g L−1), KH2PO4 (2.64 g L−1),NH4Cl (3 g L−1), MgSO4 (0.24 g L−1) and 1 mLof trace element solution (200 mg (NH4)Fe(SO4)2•6H2O, 5 mg ZnSO4•7H2O, 5 mg MnCl2•4H2O,2 mg CuSO4•5H2O, 2 mg NaB4O2•10H2O, 2 mgNaMoO4•2H2O per litre in 1 mol L−1 HCl) ata pH of 7.16 Biomass was harvested during theexponential growth phase and stored in nutrient brothcontaining 20% (v/v) glycerol at −80 ◦C prior to usein biodegradation experiments.

Biodegradation of PHA filmsThe mixed microbial culture was used to inoculate100 mL of enrichment media containing 2 g glucose,2 g yeast extract and 2 g peptone per litre and thenincubated with mixing (200 rpm, 25 ◦C). This pre-culture was collected during the exponential growthphase via centrifugation (6000 × g, 10 min) and addedto minimal medium (900 mL) to attain an opticaldensity reading of 0.18 at 660 nm. This inoculumculture was divided into three equal portions of300 mL and added to each of three 3 L sealable, sterilecontainers. Solvent cast films of PHB (×14), P(HB-co-8HV) (×14) and PHO (×14) were added to eachcontainer, were subsequently incubated for 10 days

at 25 ◦C and then mixed gently through rotation(200 rpm) under sterile air flow. Three separate sterilecontrols were used. Abiotic degradation of the polymerfilms was investigated using sterile media containingantibiotics (300 mg spectinomycin, 30 mg polymyxinB, 60 mg kanamycin per litre). A separate experimentwas conducted where polymer films were removed ona daily basis and gently vortexed in 0.85% NaCl toremove loosely attached cells. All experimental workwas conducted using aseptic techniques.

Measuring planar coverage of stronglyand loosely attached cellsDuplicate test samples of PHA films were periodicallyremoved from each of the three containers. Polymerfilms were vortexed for 5 s in 0.85% NaCl to removeloosely attached cells. Loosely attached cells werecentrifuged at 5000 × g for 5 min, washed in water,centrifuged again and dried at 80 ◦C for 2 daysbefore their dry weight was determined. Similarly,the dry weight of free-floating cells in media wasmeasured after removing three 1 mL aliquots fromeach experiment at each time point, centrifuging,washing and drying.

Polymer films with cells strongly attached werestained with SYTO 9 nucleic acid stain (MolecularProbe Inc., Eugene, USA) for imaging of remainingbiofilms on the polymer surface using a CLSMinstrument (Leica TCS SP Confocal DMIRB,Germany) equipped with an argon laser (excitation488 nm, emission wavelength 520–550 nm). Multipleimages of the biofilm and polymer surfaces wererecorded through the z plane (step size = 0.5 µm).These images were subsequently analysed using AdobePhotoshop, and the percentage of film surface areacovered by biofilm was determined by countingpixels.19 A mean surface coverage was determinedfrom the average of at least 10 images.

Measurement of total biofilmDuplicate polymer films were periodically removedfrom the in vitro biodegradation containers and theirbiofilms fixed by heating at 80 ◦C for 20 min. Thepolymer and associated biofilms were then stainedwith crystal violet using a method modified fromNarisawa et al.20 Briefly, polymer films were soakedin a 0.1% (w/v) solution of crystal violet for 60 minand then washed with minimal medium until no morestain could be removed. The crystal violet boundto the biofilm was subsequently removed by elutingwith 2 mL of 95% ethanol. The absorbance of thiscrystal violet solution was measured at 590 nm, andcorrected for background. To determine backgroundabsorbance, undegraded polymer pieces (×3) of thesame size and weight were subjected to the samestaining process.

Polymer film characterisationBiofilms were removed by soaking the polymer filmsin a solution containing 0.25% sodium hypochlo-rite, 0.1% Tween-85 and 0.01% Savinase Ultra

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(Novozyme, Australia) for 2 h, followed by ultrasoni-cation for 20 min. The polymer films were then stainedwith Nile Blue and the surfaces imaged with multiplesections through the z plane using CLSM (excita-tion 543 nm, emission 550–645 nm). For a separateseries of films at comparable times, removal of thebiofilm was followed by drying in a vacuum desiccatorfor 48 h then standing until they attained a constantweight. These weights were compared to their unde-graded counterparts and the percentage degradationcalculated. Sterile polymer surfaces were imaged withCLSM in reflection mode (excitation 458 nm, emis-sion 440–470 nm). Multiple images through the zplane (step size = 0.5 µm) were recorded. The averagesurface roughness values (Ra) were calculated fromthese images using ImageJ software (National Insti-tutes of Health, USA) according to ISO 4298 (2000):

Ra = 1/L∫ L

0|z| dx (1)

where L is the sampling length, z the plane and dxthe variations of irregularities from the mean line. Tenimages were taken and the average Ra and mean facetorientation (MFO) values determined.23

The surface hydrophobicities of undegraded poly-mer films were measured with a Rame Hart USANRL contact angle goniometer (model 100-00). A3 µL drop of Milli-Q water was placed onto the poly-mer film surfaces and an average of 6 readings foreach side recorded. These readings were then used todetermine the average surface hydrophobicity. Mediaabsorption by the polymer films was measured afterthe samples were removed from the sterile media at9 days. Absorption was measured by the equation

Absorption = Wet weight − dry weightWet weight

× 100%

(2)

The wet weight was obtained by gently blottingeach side of the polymer on absorbent paper priorto gravimetric analysis. A total of 5 pieces of eachpolymer film were used to determine the mean mediaabsorption.

Undegraded solvent cast PHA films were coatedwith gold before visualising with SEM. Partiallydegraded polymer films had their biofilms removed(as described above) and dried prior to becominggold coated. Polymer films were examined atmagnifications of ×1000, ×2500 and ×5000 at10.0 kV with a spot size of 4.0, with an FEI Quanta200 electron microscope (Brunswick, USA).

RESULTS AND DISCUSSIONPolymer biodegradation and biofilm formationSolvent cast films of PHB, P(HB-co-8HV) and PHOwere incubated in the mixed microbial culture andsamples periodically removed. In all experiments,

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Figure 1. Biofouling and biodegradation of PHA films, monitored bybiofilm coverage and polymer residual weight. (a) Residual weight ofPHB (�) and P(HB-co-8HV) (�) films, biofilm coverage of PHB (♦) andP(HB-co-8HV) (�) films. (b) Regression analysis between biofilmcoverage and biodegradation of PHB (°) and P(HB-co-8HV) (ž)solvent cast films.

measurable weight loss of PHB and P(HB-co-8HV)occurred within 10 days, and was accompaniedby a substantial biofilm formation. In contrast,no weight loss of PHO was observed. Crystalviolet staining of the cells attached to surfaces ofPHB and P(HB-co-8HV) was used to quantify thetotal biofilm on these degradable polymer surfaces.A relationship between biofouling and PHA filmdegradation, as monitored by weight loss, wasobserved (Fig. 1(a)). As the PHA films degraded, therewas a corresponding increase in the attached biofilm.A statistically significant relationship (R2 = 0.96)between biopolymer weight loss and biofilm coveragewas determined, highlighting the importance ofbiofilm formation in PHA biodegradation (Fig. 1(b)).Furthermore, this relationship suggests the potentialof predicting material loss based on the initialbiofilm coverage, in environments populated by highconcentrations of PHA degraders. The technique alsodemonstrated that in this series of experiments, filmsfabricated from P(HB-co-8HV) had apparently morebiofilm than that of their PHB counterparts, at similar

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residual weights. Analysis of surface rugosity revealeda higher original surface rugosity for PHB than forP(HB-co-8HV), suggesting that other factors, such ashydrophobicity, affected biofilm attachment to thesepolymers.

Biofilm composition and biodegradationFluorescent staining was applied to the PHA films tovisualise their attached biomass and clearly showed agradual increase in microbial attachment to polymerfilm surfaces with duration of incubation in thebiodegradation model (Fig. 2). Figures 2(a)–(c) showthe increase in microbial adhesion for a PHB film after2, 4 and 6 days. In contrast, microbial attachmentto films of P(HB-co-8HV) at the same time periodsappeared greater (Figs 2(d)–(f)).

In a separate experiment, biofouling of the PHAfilm surfaces was analysed in terms of cellularattachment. Three aliquots of media were periodicallyremoved from the in vitro degradation experimentand gently centrifuged to measure free-floating cellsnot associated with the biofilm, termed ‘planktoniccells’. The polymer film was then vortexed to removethe ‘loosely attached’ cell mass, while the residualpolymer-associated biomass was defined as ‘strongly

(a)

(b)

(c)

(d)

(e)

(f)

Figure 2. Fluorescence micrographs (100 × 100 µm) illustratingbiofouling of PHA films after 2, 4 and 6 days incubation in an in vitrobiodegradation model: (a–c) PHB; (d–f) P(HB-co-8HV).

attached’. According to this classification, changesin biofilm development and the corresponding PHAmaterial loss were quantified.

Fluorescent staining and analysis with CLSMshowed that after 2 days of incubation in thedegradation in vitro model, approximately 10% ofPHB and P(HB-co-8HV) films were covered withstrongly attached biomass corresponding to negligibleweight loss (Figs 3(a) and (b)). Biofouling of the scl-PHA films increased with length of incubation andwas accompanied by biodegradation of the films, asmonitored by gravimetry (Figs 3(a) and (b)). After9 days incubation in the accelerated biodegradationmodel, approximately 80% of the PHB film’s surfaceareas were covered with cells (Fig. 3(a)). This increasein surface coverage corresponded to a 40% weightloss (Fig. 3(a)). Qualitative observations suggestedthat the number of loosely attached cells exceededthose strongly attached to the polymer; these cellscould be readily observed as ‘clumps’ whereas CLSMwas required to visualise the strongly attached cells(Fig. 2). In support of these observations, the dryweight of the loosely attached biomass per polymerfilm rose dramatically after 4 days of incubation from1.9 to 109 mg at day 9 (Fig. 3(a)). Similar patterns ofbiofouling and biodegradation were also determinedfor solvent cast films of P(HB-co-8HV) (Fig. 3(b)).Regression analysis of the data revealed a non-significant relationship between PHA biodegradationand the strongly attached biomass (as measured bysurface area covered; R2 = 0.78). In contrast, therewas a strong correlation between PHA biodegradationand the loosely attached cells (as measured by biomass;R2 = 0.95). Thus, our results suggest that the looselyattached cells, by virtue of their numbers, might playa larger role in PHA biodegradation than previouslyobserved through electron microscopy to be stronglyattached to the polymer films.16,17

In a separate experiment, coverage of the scl-PHAfilms by strongly attached biomass was maintainedwhile the weight of loosely attached cells wassignificantly reduced by 82% to 20 mg after 9 daysof incubation (Figs 3(c) and (d)). Consistent withthe results of a regression analysis, this comparativereduction in the loosely attached biomass wasaccompanied by a significant reduction in thedegradation of the PHA films, which showedapproximately 10% weight loss, compared to 50%when the loosely attached biomass was approximately5 times greater (Fig. 3(b) versus Fig. 3(d)). This wouldappear to support the suggestion that the looselyattached microbial cells are primarily responsible forscl-PHA and may do this by virtue of extracellularenzymes.

Microbial attachment to the scl-PHA films waspredominantly in the form of flat undifferentiatedbiofilms. Similar loosely attached biofilms lackingmicrocolonies have been observed in drinking watersystems and in nutrient-poor conditions for certain

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Figure 3. Biofouling and biodegradation of (a, c) PHB and (b, d) P(HB-co-8HV) solvent cast films; residual weight of PHA (�), cells stronglyattached to films (♦), cells loosely attached (�), planktonic microbial cells (ž). In vitro biodegradation models with enhanced microbial content(a, b) and with a reduced concentration of loosely attached cells (c, d).

species such as Serratia marcescens.24,25 The influ-ence of species type in biofilm formation has beendemonstrated by investigations on biofilm morpholo-gies of three Pseudomonas species.26 Under the samenutrient conditions, P. aeruginosa formed flat undif-ferentiated biofilms, P. aureofaciens tended to formmicrocolonies and P. fluorescens formed an intermedi-ate phenotype.26 The flat type of biofilm attached tothe scl-PHA films observed here presents the greatestarea of bacteria in contact with the polymer films.Using SEM, Nishida and Tokiwa have suggested thatsmall ‘pits’ observed on the surface of PHB films weredue to such microbial colonisation.23 In our studies,however, biopolymer films were physically marked byscoring the polymer surface and this location was usedto compare the precise location on the polymer sur-face with and without the presence of biofilm. After9 days of incubation in the in vitro degradation modelboth PHB and P(HB-co-8HV) films were covered withextensive biofilms (Figs 3(a) and (b)). Removal of thisbiofilm and alignment with the CLSM images revealedthat no relationship between microcolonies and sur-face pitting could be discerned. This may be due tothe presence of flat, undifferentiated biofilms ratherthan microcolonies.

Under the experimental conditions used here, thesolvent cast PHB and P(HB-co-8HV) films showed50% of their weight loss after approximately 9.7 and8.8 days, respectively (t50). Consistent with previousreports, the more amorphous scl-PHA copolymerdegraded more quickly than the more crystallinehomopolymer. Sterile controls containing antibioticswere conducted to monitor abiotic hydrolysis. Underthese conditions, the PHB films showed 0.54 ± 0.05%weight loss and the P(HB-co-8HV) films 0.17 ±0.05% after 10 days of incubation, demonstrating thatthe majority of weight loss in the biodegradationexperiments was due to biotic rather than abioticfactors. This comparatively negligible rate of abiotichydrolysis is similar to that reported by Doi et al.1 fortemperatures under 28 ◦C.

Based on our results we can suggest that the morenumerous loosely associated cells in the biofilm playa larger role in biodegradation of the biopolymerthan their strongly attached counterparts. Thus, thebacteria do not need to be strongly adhered to thepolymer in order to degrade it.27 These results mayhave implications in devising anti-fouling strategies,which usually focus on prevention of strongly attachedmicrobial colonisation.28,29

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Figure 4. Biofouling and biodegradation of PHO: residual weight (ž)and biofilm (°) coverage. Fluorescence micrograph illustratingminimal biofouling (ca 5%) of PHO film after 6 days incubation in thein vitro biodegradation model (scale bar = 20 µm).

The biodegradability of PHO has been previouslydemonstrated in vivo and by isolates from soil;however, in the in vitro biodegradation experimentsdescribed here, PHO did not degrade.6,7,30 Consistentwith the trends exhibited by the scl-PHA, thislack of biodegradation for the mcl-PHA film wasaccompanied by negligible biofilm coverage, whichremained constant at approximately 5% for theduration of the experiments (Fig. 4). The lack ofPHO biodegradation may be due to the absence ofappropriate isolates, a consequence of the glucoseenrichment procedure. However, Tan and co-workershave shown that environmental degradation of PHOin soil was not as great as that of PHB.6,7

Surface rugositySEM of PHB and P(HB-co-8HV) films prior to andduring degradation showed a gradual increase in theroughness of the film surfaces (Figs 5(a)–(d)). After50% material weight loss, the films of PHB visuallyappear to be rougher than those of the copolymercounterpart (Figs 5(b) and (d)). In contrast, SEMimages of PHO showed no significant surfaceerosion due to in vitro biodegradation, consistent withthe lack of weight loss (Figs 5(e) and (f)). Thequalitative SEM observations were supported by three-dimensional mapping of the polymer film surfacesusing Nile Blue staining and analysis with CLSM.The microtopographies shown in Fig. 6 clearly showthe increase in surface rugosity for PHB and P(HB-co-8HV) films as a consequence of their biodegradation(Figs 6(a)–(d)). In contrast, there was no significantchange in PHO film surfaces (Figs 6(e) and (f)).Such qualitative observations are limited in relatingthe changes in surface rugosity to biofouling andbiodegradation. Consequently, changes in surfacerugosity of the polymer films due to degradation werealso monitored quantitatively through their ‘averageroughness’ (Ra).

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Figure 5. SEM micrographs of undegraded (a) PHB,(c) P(HB-co-8HV) and (e) PHO solvent cast films; surfaces of (b) PHBand (d) P(HB-co-8HV) films after 50% weight loss in the in vitrobiodegradation model; (f) PHO films demonstrated negligible weightreduction (scale bar = 50 µm).

Average roughness is defined as ‘the averagedeparture of the surface from the mean surfaceplane at a given moment’ (ISO4287:1997). Theundegraded PHB, P(HB-co-8HV) and PHO filmshad Ra values of 1.14 ± 0.18, 0.72 ± 0.15 and1.50 ± 0.2 µm, respectively. After 50% weight loss,the Ra value of the PHB film increased to 1.72 µm,corresponding to a roughness ratio change from 1:1 to1:1.5 ± 0.2. The Ra values for the films of P(HB-co-8HV) exhibiting 50% biodegradation increased by0.6 µm, a roughness ratio change to 1:1.76 ± 0.3(Fig. 7). The Ra value for the PHO films remainedunchanged. The biodegradation-induced increase insurface roughness for the scl-PHAs shown hereis consistent with reports by Ashby et al.,31 whodescribed an increase in the surface irregularity andsurface area of PHB after biodegradation by individualPseudomonas and Comamonas species.

The surface rugosity of the PHA films wasanalysed further for structures under 5 µm, definedas ‘microroughness’, and structures larger than 5 µm,termed ‘waviness’ (ISO4287:1997). In all PHA filmsexamined, the increases in surface roughness werepredominantly due to increases in the ‘waviness’component (Fig. 7). During biodegradation, themicroroughness of PHB increased slightly to 1.12 ±0.19 times its original, while the waviness increased

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Figure 6. Microtopographies of undegraded (a) PHB, (c) P(HB-co-8HV) and (e) PHO solvent cast films; microtopographies of (b) PHB and(d) P(HB-co-8HV) films after 50% weight loss in the in vitro biodegradation model; (f) PHO films demonstrated negligible weight reduction.

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4

0 100 10

Rou

ghne

ss r

atio

Time (day)

(a)

(c) (d)

(b)

5 5

Figure 7. Change in roughness ratios of solvent cast PHA films with exposure in the in vitro biodegradation model and (a, b) the sterile PHBcounterpart and (c, d) P(HB-co-8HV); overall roughness (°), microroughness (�) and waviness (�).

Polym Int 57:1042–1051 (2008) 1049DOI: 10.1002/pi

CA Woolnough et al.

by a factor of 1.72 ± 0.26 (Fig. 7(a)). A similar1.16 ± 0.33-fold increase in microroughness was alsoobserved for the P(HB-co-8HV) films; however, thewaviness component increased significantly by a factorof 3.69 ± 0.33 (Fig. 7(c)). Changes in surface rugositydue to hydrolytic degradation were less (Figs 7(b) and(d)). Further studies are required in order to relatesuch rugosity changes to film crystalline structure andthe established trend of preferential degradation of theamorphous regions.32

Individual microbial cells attached to the PHA filmswere approximately 2 µm in length (Fig. 2). Thus,polymer erosion by strongly attached cells would resultin an increase in microroughness structures (<5 µm).Similarly, an increase in the polymer surface wavinesscomponent may be due to erosion by numerouscells close together as in an undifferentiated flatbiofilm. Thus, the comparatively greater increasein waviness is consistent with the suggestion thatbiodegradation is primarily due to the relatively highconcentrations of loosely attached cells in the biofilm(Fig. 3).

Changes in the average angle of facets in reference tothe horizontal plane can be an important contributorto the type of surface roughness and is directlyrelated to surface gloss; the smaller the MFO, themore glossy the surface.33 The MFO of the scl-PHAfilms was measured before and after biodegradation(Fig. 8). After biodegradation, there was an increasein the MFO for both polymers, +2.7◦ and +3.6◦for PHB and P(HB-co-8HV), respectively, indicatingthat both polymers had become less glossy (Fig. 8).Yasin and Foster, reporting the in vivo environmentaldegradation of scl-PHA films, observed similar changesin their ‘gloss factor’ results.34 Abiotic degradationcontrols contained polymer films immersed in sterilemedium for 9 days and resulted in a +0.9◦ increase inthe facet orientation for the PHB films and a −0.8◦change for their P(HB-co-8HV) counterparts (Fig. 8).Thus abiotic degradation slightly reduced the glossof the PHB films but had the opposite effect for theP(HB-co-8HV) films.

-2

-1

0

1

2

3

4

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nge

in M

ean

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rient

atio

n (°

)

H B

Figure 8. Change in mean facet orientation of surfaces of PHAsolvent cast films after 50% weight loss due to hydrolytic degradation(H) and biodegradation (B): PHB ( ) and P(HB-co-8HV) ( ).

Polymer surface and bulk hydrophobicityEginton et al. have reported that hydrophobicity ratherthan surface roughness influenced the strength ofbiofilm attachment to polyethylene films.15 In theexperiments reported here, mean contact angles ofthe undegraded polymer films were determined asa measure of surface hydrophobicity, with littledifference between PHB (73.3◦), P(HB-co-8HV)(75.3◦) and PHO (73◦). Choi et al.35 have reportedthat an increase in the valerate content causes anincrease in surface hydrophobicity; thus the slightlyhigher surface hydrophobicity for the P(HB-co-8HV)films used in these experiments is consistent withthis and may have facilitated cell adhesion, wherethe copolymer films displayed a more rapid coverageof strongly attached cells (Figs 2 and 3).33 However,contact angles reflect only the surface and not thebulk hydrophobicity of the polymer. Gravimetry wasused to determine the latter where the absorptionof sterile media by the polymer films showed arelatively low water uptake of 2.70 ± 0.3% for PHB,1.61 ± 0.06% for P(HB-co-8HV) and 0.75 ± 0.05%for PHO after 9 days of incubation. Water adsorptionby the polymers is consistent with the establishedmechanisms for their degradation, where penetrationby extracellular depolymerase enzymes leads to a chaincleavage and preferential erosion of amorphous regionsin the polymer.14

CONCLUSIONSPHAs are a family of biopolymers with potentialas environmentally friendly, commercial thermoplas-tics, the biodegradation of which has been studiedextensively in a variety of environments.1 Despite therecognition that microbial ecology plays a decisive rolein the environmental biodegradation of these poly-mers, there are few studies attempting to investigatethe relationship between microbial colonisation andpolymer degradation. We have used an acceleratedin vitro biodegradation model composed of an unde-fined mixed microbial culture to study this relationshipfor an scl-PHA homopolymer (PHB), P(HB-co-8HV)copolymer and an mcl-PHA terpolymer (PHO). In thisstudy, a statistically significant link was establishedbetween polymer weight loss and biofilm formation.Furthermore, our results suggest that this degradationwas primarily conducted by cells loosely attached tothe polymer rather than those strongly attached, as pre-viously suggested.19 Biofilm formation and its type aredependent upon numerous factors; the flat undifferen-tiated biofilms observed in this study were responsiblefor a gradual increase in surface rugosity, primarilyobserved as an increase in waviness. We have devel-oped protocols based on a combination of CLSM andCAG to qualitatively and quantitatively map surfacechange in relation to biofilm formation and polymerbiodegradation. We are currently using these protocolsto investigate strategies to inhibit cellular adhesion.The results reported here have implications in polymer

1050 Polym Int 57:1042–1051 (2008)DOI: 10.1002/pi

Biodegradation and biofouling of PHA films

design and surface modification for the prevention ofbiofouling in medical and environmental applications.

ACKNOWLEDGEMENTSThe authors wish to thank Dr Halasz for CLSMsupport. C. Woolnough was funded by an AustralianPostgraduate Award.

REFERENCES1 Doi Y, Kanesawa Y, Kunioka M and Saito T, Macromolecules

23:26 (1990).2 Mergaert J, Webb A, Anderson C, Wouters A and Swings J,

Appl Environ Microbiol 59:3233 (1993).3 Brandl HGR, Lenz RW and Fuller RC, Adv Biochem Eng

Biotechnol. 41:77 (1990).4 Steinbuchel A and Valentin HE, Fems Microbiol Lett 128:219

(1995).5 Brandl H, Gross RA, Lenz RW and Fuller RC, Advances in

Biochemical Engineering/Biotechnology. Springer, Berlin (1990).6 Ho YH, Gan SN and Tan IK, Appl Biochem Biotechnol

102–103:337 (2002).7 Lim SP, Gan SN and Tan IKP, Appl Biochem Biotechnol 126:23

(2005).8 Abe H and Doi Y, Int J Biol Macromol 25:185 (1999).9 Mergaert J, Wouters A, Anderson C and Swings J, Canad J

Microbiol 41:154 (1995).10 Bonhomme S, Cuer A, Delort AM, Lemaire J, Sancelme M and

Scott G, Polym Degrad Stab 81:441 (2003).11 Linos A, Berekaa MM, Reichelt R, Keller U, Schmitt J, Flem-

ming HC, et al, Appl Environ Microbiol 66:1639 (2000).12 Maeyama R, Kwon IK, Mizunoe Y, Anderson JM, Tanaka M

and Matsuda T, J Biomed Mater Res 75A:146 (2005).13 Fox P, Suidan MT and Bandy JT, Water Res 24:827 (1990).14 Pasmore M, Todd P, Smith S, Baker D, Silverstein J, Coons D,

et al, J Membr Sci 194:15 (2001).

15 Eginton PJ, Gibson H, Holah J, Handley PS and Gilbert P,Colloids Surf B: Biointerfaces 5:153 (1995).

16 Mabrouk MM and Sabry SA, Microbiol Res 156:323 (2001).17 Molitoris HP, Moss ST, de Koning GJM and Jendrossek D,

Appl Microbiol Biotechnol 46:570 (1996).18 Foster LJR and Tighe BJ, Polym Degrad Stab 87:1 (2005).19 Weitere M, Bergfeld T, Rice SA, Matz C and Kjelleberg S,

Environ Microbiol 7:1593 (2005).20 Narisawa N, Furukawa S, Ogihara H and Yamasaki M, J Biosci

Bioeng 99:78 (2005).21 Foster LJR, Russell RA, Sanguanchaipaiwong V, Stone DJM,

Hook JM and Holden PJ, Biomacromolecules 7:1344 (2006).22 Yu J, Biofouling 19:83 (2003).23 Nishida H and Tokiwa Y, J Environ Polym Degrad 3:187 (1995).24 Martiny AC, Jorgensen TM, Albrechtsen HJ, Arvin E and

Molin S, Appl Environ Microbiol 69:6899 (2003).25 Rice SA, Koh KS, Queck SY, Labatte M, Lam KW and

Kjelleberg S, J Bacteriol. 187:3477 (2005).26 Heydorn A, Nielsen AT, Hentzer M, Sternberg C, Givskov M,

Ersboll BK, et al, Microbiol UK 146:2395 (2000).27 Foster LJR, Lenz RW and Fuller RC, in Hydrogels and

Biodegradable Polymers for Bioapplications, ed. by Ottenbrite R,Park K and Huang S. ACS Books, Washington, DC,p. 68 (1996).

28 Foster LJR, Sanguanchaipaiwong V, Gabelish C and Hook J,Polymer 46:6587 (2005).

29 Hany R, Bohlen C, Geiger T, Schmid M and Zinn M,Biomacromolecules 5:1452 (2004).

30 Schirmer A, Jendrossek D and Schlegel HG, Appl EnvironMicrobiol 59:1220 (1993).

31 Ashby RD, Cooke P and Solaiman DKY, J Polym Environ15:179 (2007).

32 Abe H and Doi Y, Macromolecules 31:1791 (1998).33 Chinga G, Stoen T and Gregersen OW, J Pulp Paper Sci 30:307

(2004).34 Yasin M and Foster LJR, Polymat ’94:162:(1994).35 Choi GG, Kim HW and Rhee YH, J Microbiol 42:346 (2004).

Polym Int 57:1042–1051 (2008) 1051DOI: 10.1002/pi