rapid, sensitive and direct analysis of exopolysaccharides from biofilm on aluminum surfaces exposed...

Research Article

Received: 3 August 2011 Revised: 21 September 2011 Accepted: 4 October 2011 Published online in Wiley Online Library

(wileyonlinelibrary.com) DOI 10.1002/jms.2003

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Rapid, sensitive and direct analysis ofexopolysaccharides from biofilm on aluminumsurfaces exposed to sea water usingMALDI-TOF MSNazim Hasan,a Judy Gopala,b and Hui-Fen Wua,b,c*

Biofilm studies have extensive significance since their results can provide insights into the behavior of bacteria on materialsurfaces when exposed to natural water. This is the first attempt of using matrix-assisted laser desorption/ionization-mass
spectrometry (MALDI-MS) for detecting the polysaccharides formed in a complex biofilm consisting of a mixed consortiumof marine microbes. MALDI-MS has been applied to directly analyze exopolysaccharides (EPS) in the biofilm formed onaluminum surfaces exposed to seawater. The optimal conditions for MALDI-MS applied to EPS analysis of biofilm have beendescribed. In addition, microbiologically influenced corrosion of aluminum exposed to sea water by a marine fungus was alsoobserved and the fungus identity established using MALDI-MS analysis of EPS. Rapid, sensitive and direct MALDI-MS analysison biofilm would dramatically speed up and provide new insights into biofilm studies due to its excellent advantages such assimplicity, high sensitivity, high selectivity and high speed. This study introduces a novel, fast, sensitive and selective platformfor biofilm study from natural water without the need of tedious culturing steps or complicated sample pretreatmentprocedures. Copyright © 2011 John Wiley & Sons, Ltd.
Supporting information may be found in the online version of this article.

Keywords: biofilm; mass spectrometry; aluminum; corrosion; EPS; marine; fungus

* Correspondence to: Hui-Fen Wu, Department of Chemistry, National SunYat-Sen University, Kaohsiung, 70, Lien-Hai Road, Kaohsiung, 80424, Taiwan.E-mail: [email protected]

a Department of Chemistry, National Sun Yat-Sen University, Kaohsiung, 70,Lien-Hai Road, Kaohsiung, 80424, Taiwan

b Center for Nanoscience and Nanotechnology, National Sun Yat-Sen University,70, Lien-Hai Road, Kaohsiung, 80424, Taiwan

c Doctoral Degree Program in Marine Biotechnology, National Sun Yat - SenUniversity, Kaohsiung, 80424, Taiwan

INTRODUCTION

Most marine environments contain only dilute substances thatcan be used for metabolism and growth. In contrast, naturalsurfaces introduced into aquatic habitats tend to collect and con-centrate nutrients by charge–charge or hydrophobic interactions [1].Bacterial colonization on abiotic materials such as suspended parti-cles, metal surfaces and concrete or on biotic surfaces is believedto be one of the microbial survival strategies because it providesthe microorganisms with important advantages, including (1)increased access to nutrients, (2) protection against toxins andantibiotics, (3) maintenance of extracellular enzyme activities and(4) shelter from predation [2].For these reasons, surfaces in contact with water are rapidly

colonized by bacteria. Free-living or a planktonic mode of growthof microorganisms is usually observed in laboratory cultures.However, this growth mode is infrequent in the natural environ-ment, and bacteria may seek out advantageous niches and thusare described by the term ‘benthic’ [1]. Once planktonic cells meetsuch surfaces, they attach and eventually develop biofilmsthrough growth and division [3,4]. The main ‘cement’ for all thesecells and products is the mixture of polysaccharides secreted bythe cells established within the biofilm. The biofilm thus consistsof two entities, microbial cells and polysaccharides. The majorcomponent in the biofilm matrix is water� up to 97% [3], andthe characteristics of the solvent are determined by the solutesdissolved in it. The exact structure of any biofilm is probably aunique feature of the environment in which it develops.The exopolysaccharides (EPS) that constitute the other major

J. Mass. Spectrom. 2011, 46, 1160–1167

component in the biofilm give the structural framework to thebiofilm. The EPS synthesized by microbial cells vary greatly intheir composition and hence in their chemical and physicalproperties. Some are neutral macromolecules, but the majorityare polyanionic due to the presence of either uronic acids(d-glucuronic acid being the commonest, although d-galacturonicand d-mannuronic acids are also found) or ketal-linked pyruvate.Inorganic residues, such as phosphate or rarely sulphate, may alsoconfer polyanionic status [5,6]. The study of biofilms hasskyrocketed in recent years due to increased awareness of thepervasiveness and impact of biofilms on natural and industrialsystems, as well as human health. In some respects, biofilmformation applies to human beings in the remediation process ofwastewater, degradation of recalcitrant, aquaculture, etc. In otherrespects, biofilm processed on heat exchangers, pipelines, shipsurfaces and other industrial devices causes serious problemsand consumes large amounts of time and money to remove

Copyright © 2011 John Wiley & Sons, Ltd.

Direct analysis of biofilm using MALDI-TOF MS

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it. Biofilms cost literally billions of dollars every year in energylosses, equipment damage, product contamination and medi-cal infections. Also, biofilm formed on implanted materials hasbeen related to microbial diseases [7,8]. Therefore, control ofbiofilm formation has been an important topic of interest todate. The complexity of biofilm activity and behavior requiresresearch contributions from many disciplines such as bio-chemistry, engineering, mathematics and microbiology. There-fore, a proper understanding of natural biofilms is crucial foreffecting control of biofilms as well as for tapping the poten-tial of biofilms for beneficial purposes.

Detailed studies have been conducted in analyzing biofilmformation in seawater [9–11]; most of these studies involved thecharacterization of the microbes occurring in the biofilm. Theobjective of the present paper is to characterize the EPS formedon Al surfaces exposed to sea water. The most widely practisedmethod for estimation of EPS is through spectrophotometricestimation of proteins and carbohydrates, respectively. Usingmatrix-assisted laser desorption/ionization (MALDI), it is possibleto obtain polysaccharide peaks which would eventually lead todetection of polysaccharides in the biofilm. MALDI is thus a moresuperior technique from this perspective and also since the bio-film was directly analyzed, which definitely is time saving. Effortshave been made to prove the use of MALDI-time-of-flight (TOF)mass spectrometry (MS) as an efficient technique in analyzingbiofilm, especially the EPS portion of the biofilm. Since this paperproposes for the first time about the use of MALDI-MS for rapidand direct analysis of biofilm from sea water, the sample prepara-tion conditions, the matrix used and the MALDI-MS ion modes(positive or negative ions) to study the EPS have been optimized.Finally, a case of biocorrosion phenomenon was observed from amarine fungus which is also reported in the present study.

EXPERIMENTAL SECTION

Chemicals and methods

2, 5-Dihydroxybenzoic acid (DHB, 99%), a-Cyano-4-hydroxycin-namic acid (CHCA, 99 %) and Trifluoroacetic acid (TFA, 99%) werepurchased from Sigma Chemical Co. (St. Louis, MO, USA). 3,5-Dimethoxy-4-hydroxycinnamic acid (SA, 98%) was purchasedfrom Alpha Aesar (UK). Acetonitrile (HPLC grade, 99.99%) andacetone (GC grade, 99.8%) were purchased from (J.T. Baker,Phillipsburg, NJ, USA). Aluminum metal sheet (0.2mm thick,99% metal basis) was purchased from Alfa Aesar (A JohnsonMatthey Company, USA), and the glass jar (500mL) was boughtfrom Schott Duran, Germany. Deionized water purified by aMilli-Q reagent system (Millipore, Milford, MA, USA) was usedfor all the experiments. Ultrasonicator (Elma, E 30 H, Elma GmbH& Co KG, Germany) was used to isolate the biofilm from the sur-face of aluminum coupons. The centrifuge (Hettich zentrifugen,Andreas Hettich GmbH & Co KG, Germany) was used for high-speed centrifugation of biofilm samples at 31 514 g, 4 �C for20min. Wide zoom camera from Apisc (model STC-C33USB,Germany) was used to photograph the macroscopic settlementsand severely corroded aluminum coupons.

Preparation of aluminum coupons for biofilm formation

Commercially pure aluminum coupons (30mm� 20mm� 0.2mm)were cut from sheets, and a hole (1mm in diameter) was madeon one side of the coupon for suspending the coupons. They wereultrasonically cleaned using soap solution for 10min and washed

J. Mass. Spectrom. 2011, 46, 1160–1167 Copyright © 2011 John

in running water and finally rinsed in deionized water and air dried.The samples and the wires used to suspend themetal coupons wereultrasonicated in acetone for 5min and then kept under ultravioletlight for surface sterilization in a laminar flow chamber for 2h.

Biofilm formation in seawater

Sea water was collected from Shiziwan Bay, Kaohsiung, Taiwan(at a distance of 6 m further into the sea from the shore). Biofilmwas formed by hanging the Al metal coupons using the sterilewires in seawater (500mL) in conical flasks (as shown in Fig. 1).The coupons were retrieved at 1 day, 2 day, 3 day, 1week, (repre-senting short-term studies), 1month and 2month intervals(representing long term studies), and then the biofilm was col-lected and studied. 200mL of seawater was removed from theconical flask and replaced with fresh seawater after 1 month.

Biofilm sample preparation prior to MS detection

The biofilm formed after each time interval was isolated by ultra-sonically cleaning the metal coupons for 15min in sterile water(3mL) [12]. The biofilm in sterile water was collected in Eppendorftubes and centrifuged for 15min (at 31 514 g, 4 �C) for EPSextraction [13]. The supernatant and precipitate formed were sep-arated, and both were analyzed by MALDI-TOF MS. Figure 1 givesthe schematic representation of the experimental setup, and alsothe methodology followed during the study.

Preparation of matrix solution

DHB (50mM), CHCA (50mM) and 3,5-Dimethoxy-4-hydroxycin-namic acid (50mM) solution were prepared in acetonitrile/deio-nized water (2:1, v/v). For analyzing high-mass polysaccharides,DHB of high concentration (161mM) was used.

MALDI-TOF MS and ESI-MS analysis

Mass spectra were obtained using a MALDI-TOF mass spectrom-eter (Microflex, Bruker Daltonics, Bremen Germany) equippedwith a nitrogen laser (337 nm), a 1.25m flight tube and thesample target having the capacity to load 96 samples simulta-neously. All spectra were acquired with the following parametersset on the MALDI-MS: IS1, 19.0 kV; IS2, 16.15 kV; lens, 9.35 kV andreflector at 20.0 kV. The laser energy was adjusted to slightlyabove the threshold to obtain good resolution and signal-to-noise ratios at 60Hz. All mass spectra were collected in boththe positive and negative ions of reflectron modes with 200 lasershots accumulated for each spectrum. Electrospray ionization(ESI)-MS for supernatant analysis was performed with an ion trapmass spectrometer (Finnigan LCQ-Advantage, Thermoquest, SanJose, CA, USA). The temperature of the capillary was set at200 �C. The electrospray voltage was maintained at 4.5 kV. Themass range was set from 50 to 2000Da. The voltages of the cap-illary and tube lens offset were set at 40 and 70 V, respectively.

RESULTS AND DISCUSSION

The EPS extracted from the biofilm was directly analyzed byMALDI-TOF MS. The detailed procedures for these biofilm studiesare described in Fig. 1. Figure 2 shows the EPS results obtainedfrom aluminum surfaces for 1, 2, 3 and 7 day (1week) intervalswhich represent the short-term studies. The results indicate thepresence of various polysaccharide peaks which increased in

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1 2 3

4 5 6 5 6 6

Initial position after insertion the coupons in sea water

After one month After two months

Glass jar (500 mL)

Sea water

Al metal coupons Wire

White deposits White deposits

Al samples with biofilm removed at different intervals of times

Coupons with biofilm immersed in 3mL sterile water and ultrasonicated for 15 minutes to remove the biofilm

The biofilm samples centrifuged at 31514 g and 4 °C for 15 minutes

Supernatant and precipitate separated and used for EPS analysis by MALDI-TOF MS

Figure 1. Schematic showing setup used for biofilm formation on aluminum surfaces and the procedures prior to MS detection.

N. Hasan, J. Gopal and H.-F. Wu

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number with exposure time up to 1 week. In contrast, the resultsfrom the long-term exposure studies (Fig. 3) reveal that the EPSpeaks did not increase steadily after 1month (Fig. 3c) and2months (Fig. 3d). Using cut-off mode on Fig. 2 and Fig. 3, wefurther show the distinct peaks obtained in the mass range of1200–1800 to 900–1800m/z in order to enhance the visibility ofthe low intensity EPS peaks as given in Fig. S1 and Fig. S2(Supporting information) respectively. The decline in these

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Figure 2. MALDI-MS spectra of EPS of biofilm formed after (a) 1 day (b) 2 d

wileyonlinelibrary.com/journal/jms Copyright © 2011 Joh

low-mass EPS peaks coincided with the increase in high-masspolysaccharide peaks which will be discussed later in this paper.The polysaccharide peaks formed on the Al surface were identi-fied using Glycobench software. GlycoWorkbench is a softwarefor semi-automatic interpretation and annotation of mass spectraof sugars. GlycoWorkbench is a suite of software tools designedfor rapid drawing of glycan structures and for assisting theprocess of structure determination from MS data. The graphical

(b)

(d)

(c)

(a)

ay (c) 3 day (d) 1week of exposure to sea water.

n Wiley & Sons, Ltd. J. Mass. Spectrom. 2011, 46, 1160–1167

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(a)

(b)

(c)

(d)

Figure 3. MALDI-MS spectra of (a) matrix only (b) sea water and EPS of biofilm formed after exposure to sea water for (c) 1month and (d) 2months. Allspectra were obtained using DHB matrix.


interface of GlycoWorkbench provides an environment in whichstructure models can be rapidly assembled, their masscomputed, their fragments automatically matched with MSndata and the results compared to assess the best candidate.GlycoWorkbench can greatly reduce the time needed for theinterpretation and annotation of mass spectra of glycans. Themass peaks obtained from the analysis of the marine biofilmwere loaded onto the glycoWorkbench software and the massvalues identified. This software is used to broadly classify thepolysaccharides into N-linked glycans and O-linked glycans.Based on this result, we have classified the polysaccharide peaksinto these two categories.

Peaks observed at m/z 531.3, 559.3, 735.2, 920.1, 925.1 wereO-linked polysaccharide moieties. N-linked polysaccharides wereobserved at m/z 735.4, 881.6, 1080.7, 1083.3, 1212.9, 1374.4,1711.9 and 1728.2. (Here, O-linked and N-linked polysaccharidewere taken from both Fig. 1 and Fig. 2). This is the first study todetect the polysaccharides formed in a complex biofilm consist-ing of a mixed consortium of microbes from exposure to seawater. Attempts were made to analyze the biofilm using ESI-MS,too, but experiments were discontinued since ESI-MS was inade-quate and insensitive for this study (Fig. S3). Thus, MALDI-TOF MSwas a sensitive and superior technique for analyzing the polysac-charides present in complex biofilms formed in the marineenvironment compared to ESI-MS.

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Detection of supernatant and precipitate from biofilm

In order to observe the difference in EPS in the supernatant(believed to contain the polysaccharides) and the precipitate(containing the bacterial cells), both supernatant and precipitatewere analyzed in MALDI-TOF MS. The results demonstrated thatdistinctly different peaks were observed in either case, and thesupernatant showed more polysaccharide peaks compared tothe bacterial cell pellet isolated by centrifugation. Figure 4provides these comparative spectra. In Fig. 4 (i), the polysaccha-ride peaks observed at m/z 412.9, 547.3 575.3, 601.4, 721.0,758.7, 796.7, 881.9, 917.7 and 974.6 were present exclusively in


the supernatant extracted from the biofilm by high-speed centri-fugation which is reported to extract the EPS from the microbialcomponents of the biofilm. The supernatant is reported tocontain the loosely bound and some of the bound EPS [14]. There-fore, it is most probable that the peaks obtained are mostly frompolysaccharides in the biofilm, although negligible peaks couldbe from peptides, too. The pellet containing the strippedmicrobial cells was also analyzed for polysaccharides. The peaksat m/z 886.6 1042.6, 1064.6, 1083.6 and 1220.5 were specificallyobserved on the cell pellets.

Effect of matrix and positive or negative ion modefor detection

The matrix effect on the EPS studies of biofilm was optimized. Dif-ferent matrices were examined, and their performance on EPSdetection was evaluated. In Fig. 5 and Fig. 6, the EPS obtainedfrom 2-month-old biofilm from the aluminum surfaces exposedto sea water was analyzed using CHCA, DHB, SA and binary ma-trix of DHB with CHCA (1:1 ratio). The results indicate that SAwas not suitable for EPS studies due to the relatively low signalsobserved. Although CHCA and the binary matrix of DHB withCHCA could obtain good signal intensity, the low-mass clusterions of these matrices were intense and caused serious interfer-ences with the EPS peaks. Among all these matrix conditions,DHB is the best choice because less background ions were ob-served at the low-mass region. In addition, DHB has also beenpreviously discussed as the most ideal matrix for polysaccharidestudies in MALDI-MS [15].

Next, attempts were made to further compare the positiveion mode with negative ion mode for EPS detection in theMALDI-MS. Conventionally, positive ion mode is the mostcommon mode for bacterial analysis due to detection of proteinions. EPS in the biofilm was also attempted in the negative ionmode since this study focused on the polysaccharide analysisrather than bacterial analysis of proteins from the biofilm. Figure 5and Figure 6 display the comparative study conducted in bothmodes. As can be seen from the spectra, the spectral pattern


3

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Figure 4. MALDI-TOF MS spectra of EPS in (i) supernatant (ii) precipitate after high-speed centrifugation of biofilm exposed to sea water for 1-month-old biofilm.


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taken in the positive ion mode and the negative ion mode areunique. Thus, a combination of both modes is required to get acomprehensive view on the polysaccharides present in thebiofilm. As we can see from Fig. 5 and Fig. 6, both the spectraacquired in positive and negative ion mode are different,although the analyte is the same (EPS). Thus, in order to get adetailed view of all the possible polysaccharides in the biofilm,it is necessary that we combine both the positive and negativeion mode. Using cutoff mode on Fig. 5 and Fig. 6 magnifies thepeaks in the range within 700–1500 to 800–1400m/z, respec-tively, to explore the low intensity EPS (Fig. S3 and Fig. S4).When operated in the positive ion mode, more peaks were

observed compared to those obtained in the negative ion mode.Although only a few peaks were observed in the case of the

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Figure 5. MALDI-TOF MS spectra of EPS from 1-month-old biofilm analyzedthe reflectron positive ion mode.


negative ion mode, it is conclusive that these peaks belong tothe polysaccharides in the biofilm. Therefore, EPS analysisconducted in negative ion mode can also provide additionalinformation for polysaccharides in the biofilm samples. Table 1provides the total list of EPS peaks observed in the marine bio-films analyzed by MALDI-MS.

Observation of microbiologically influenced corrosion

After an exposure period of 1 month, a small macroscopic depo-sition (1mm) was observed on some aluminum coupons. Thissettlement was observed to increase in size (4.0mm in diameter)and after a period of 2 months when the coupons were removedto retrieve the biofilm for EPS studies. It was observed that below

(a)

(b)

(c)

(d)

using a different matrix. (a) CHCA, (b) DHB, (c) CHCA+DHB and (d) SA in


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(c)

(b)

(a)

Figure 6. MALDI-TOF MS spectra of EPS from 1-month-old biofilm analyzed using a different matrix. (a) CHCA, (b) DHB, (c) CHCA+DHB and (d) SA inthe reflectron negative ion mode.


the macro settlement, the aluminum surface was completelycorroded, and crevice was observed (4.0mm in diameter).Figure 7a shows the photograph of the coupon with patchysettlement and signs of localized corrosion beneath it after themacroscopic settlement were removed. The EPS isolated duringthis time period indicated the presence of a high-mass EPS calledpullulan. Figure 7b shows the spectra of this high-mass EPS,having a repetitive spectral profile. Spectrum Fig. 7b (i) is fromthe fungus (white flakes) present in the seawater in which thecoupons were exposed, and spectrum Fig. 7b (ii) is obtained fromthe biofilm on the aluminum surfaces showing similar homolo-gies to the higher molecular weight EPS. Hsu et al. reportedthe existence of such a high-mass polysaccharide moiety at4000–6000Da with a similar profile which is the characteristicof these polysaccharide spectra [16].

The deterioration of metals due to microbial activity is termedbiocorrosion or microbially influenced corrosion (MIC). Owing toits economic and environmental importance, MIC has been thesubject of extensive studies for the past 5 decades. Bacteria are

Table 1. Consolidated list of exopolysaccharide peaks obtained using a dimarked in bold are those that appear even when a different matrix or mod

CHCA matrix DHB matrix

Positive ion modem/z

Negative ionmode m/z



Po

335.2 458.1 312.5 481.8

378.6 479.1 397.0 536.9

440.9 588.2 532.7 657.0

572.2 647.3 743.5 681.9

612.0 735.3 881.6 713.0

651.5 779.2 920.1 859.1

829.2 805.9 1080.7 889.9

867.0 1035.0


considered the primary colonizers of inanimate surfaces in bothnatural and man-made environments. Therefore, the majority ofMIC investigations have addressed the impact of pure or mixedculture bacterial biofilms on corrosion behavior of iron,copper, aluminum and their alloys. Biocorrosion is a result ofinteractions, which are often synergistic, between the metal sur-face, abiotic corrosion products, and bacterial cells and theirmetabolites. The metabolites include organic and inorganic acidsand volatile compounds, such as ammonia and hydrogen sulfide.Biocorrosion produced from fungi is typically not common, but ifpresent, it is more severe, since fungi secrete acidic metaboliteswhich corrode the metals. Also, their bigger size and spreadinghyphae could accelerate corrosion compared to the microscopicbacterial corrosion [17].

The white macroscopic growth observed on the aluminumsurface was removed and spotted on a MALDI target plate andanalyzed using a DHB matrix. The same polysaccharide peakswere obtained from this organism too (Fig. 7b (i)). The polysac-charides identified as pullulan from literature [16] are produced

fferent matrix and also positive and negative ion modes. The m/z valuese was used

CHCA+DHB matrix SA matrix

sitive ion modem/z




375.1 481.8 320.0 446.6

398.1 573.3 372.1 670.9

532.1 657.0 397.1 693.1

829.3 681.9 472.1 917.5

1058.6 713.1 487.1 1141.9

779.1 509.0 1164.2

859.1

889.9

1035.0

1065.6

1211.2


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Microscopic microbial settlement Severe corroded and crevice part of Al coupon

(i) (ii)

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(ii)

(a)

(b)

Figure 7. a. Photograph of aluminum coupons retrieved after an exposure period of 2 months from sea water showing (i) macroscopic microbial set-tlement (ii) severely localized corrosion below the microbial attachment. b. MALDI-TOF spectra of high-mass polysaccharides (pullulan) observed on the(i) fungus (white flakes) present in the seawater in which the coupons were exposed (ii) biofilm on the aluminum surfaces showing similar homologies.


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by a specific fungus called Aureobasidium pullulans. This fungus isreported to be present in seawater and is also mentioned to be acosmopolitan fungus, so it is not unusual, with respect to its habitatthat it was present in the biofilm on aluminum surfaces exposed toseawater. The significance of this fungus (Aureobasidium pullulans)is that it can degrade plastics and produces a lot of enzymesand proteases [18]. Also, Srivatsava et al. reported the incidenceof microbiologically influenced corrosion by this fungus onaluminum exposed to aircraft fuels. They reported that alumi-num is highly susceptible to form microbiologically influencedcorrosion in the presence of this fungus [19]. There is also a directimplication of the production of high molecular polysaccharideby this fungus which is pullulan. Pullulan, an extracellular water-soluble polysaccharide produced by Aureobasidiumpullulans, is a linear a-D-glucan, consisting mainly of maltotrioseunits interconnected by a (1! 6) linkage [20]. Pullulan producesa high-viscosity solution at a relatively low concentration andcan be used for oxygen impermeable films, thickening orextending agents or adhesives [21]. Thus, pullulan due to itsadhesive property can enhance biofilm adhesion and owing toits oxygen impermeability facilitate differential aeration cells,depriving the surface of oxygen, disrupting the integrity of theprotective oxide film, leading to severe localized biocorrosion.


Thus, Aureobascidium pullulans is a good model for studyingbiocorrosion of aluminum, and it has high implication to MICof aluminum.

CONCLUSION

The study first demonstrated that MALDI-TOF MS can be appliedas a rapid, sensitive, selective and powerful technique for directlyanalyzing exopolysaccharides in complex biofilms formed in seawater. The ideal matrix for the biofilm studies was DHB, and thestudy also emphasizes the use of negative ion detection to gainmore information for EPS in addition to the positive ion detectionfor characterizing the EPS in the biofilm. A predominant funguswas observed in the biofilm, and a series of high-mass polysaccha-ride peaks were also obtained corresponding to pullulan. Thealuminum surfaces below the fungus showed severely localizedcorrosion. This MIC was observed after an exposure period of1month, indicating that aluminum is susceptible to form MIC inthe presence of the fungus Aureobascidium pullulans. The currentapproach establishes a rapid and sensitive approach using MALDI-TOF MS for direct detection of EPS in biofilm without culturing orseparation pretreatments.



Acknowledgements

We thank the National Science Council (Taiwan) for the financialsupport.

Supporting Information

Supporting information may be found in the online version ofthis article.

REFERENCES[1] T. J. Beveridge, S. A. Makin, J. L. Kadurugamuwa, Z. Li. Interactions

between biofilms and the environment. FEMS Microbiol. Rev. 1997,20, 291.

[2] H. Dang, C. R. Lovell. Bacterial Primary Colonization and Early Succes-sion on Surfaces in Marine Waters as Determined by Amplified rRNAGene Restriction Analysis and Sequence Analysis of 16S rRNA Genes.Appl. Environ. Microbiol. 2000, 66, 467.

[3] Z. Xiaoqi, P. L. Bishop, M. J. Kupferle. Measurement of polysaccharidesand proteins in biofilm extracellular polymers. Water Sci. Technol.1998, 37, 345.

[4] K. K. Kwon, H. S. Lee, S. Y. Jung, J. H. Yim, J. H. Lee, H. K. Lee. Isolationand Identification of Biofilm-Forming Marine Bacteria on GlassSurface in Dae-Ho Dike Korea. J. Microbiol. 2002, 40, 260.

[5] I. W. Sutherland. Microbial exopolysaccharides-structural subtletiesand their consequences. Pure Appl. Chem. 1997, 69, 1911.

[6] I. W. Sutherland. Biofilm exopolysaccharides: a strong and stickyFramework. Microbiology 2001, 147, 3.

[7] J. W. Costerton, K. J. Cheng, G. G. Geesey, T. I. Ladd, J. C. Nickel, M.Dasgupta, T. Marrie. Bacterial biofilms in nature and disease. Ann.Rev. Microbiol. 1987, 41, 435.

[8] F. R. Burton, F. Correia, J. M. DiRienzo. Corncobs: a model for oral micro-bial biofilms, H. J. Busscher, L. V. Evans, Editors, Oral Biofilms and PlaqueControl, Harwood Academic Publishers: Amsterdam, 1998, 145–162.

[9] V. Thiyagarajan, V. P. Venugopalan, K. V. K. Nair, T. Subramoniam.Macrofouling in the cooling conduits of Madras Atomic PowerStation. Ind. J. Mar. Sci. 1997, 26, 305.


[10] S. Rajagopal, K.V.K. Nair, A. Azariah. Response of brown mussel, Pernaindica to elevated temperature in relation to power plant biofoulingcontrol. J. Therm. Biology. 1995, 20, 461.

[11] J. Gopal, P. Muraleedharan, H. Sarvamangala, R. P. George, Dayal RK,B. V. R. Tata, H. S. Khatak, K. A. Natarajan. Biomeneralisation of manganeseon titanium surfaces exposed to seawater. Biofouling 2008, 24, 275.

[12] J. Gopal, R. P. George, P. Muraleedharan, H.S. Khatak. Photocatalyticinhibition of microbial adhesion by anodized titanium. Biofouling2004, 20, 167.

[13] X. Pan, J. Liu, D. Zhang, C.X. Xi, L. Li1, W. Song, J. Yang, A comparisonof five extraction methods for extracellular polymeric substances(EPS) from bio-film by using three dimensional excitation-emissionmatrix (3 DEEM) fluorescence spectroscopy. Water Sur. Anal. 2010,36, 111–116.

[14] X. Y. Li, S. F. Yang. Influence of loosely bound extracellular polymericsubstances (EPS) on the flocculation, sedimentation and dewater-ability of activated sludge. Water Res. 2007, 41, 1022.

[15] B. Stahl, M. Steup, M. Karas, F. Hillenkamp. Analysis of neutral oligo-saccharides by matrix-assisted laser desorption ionization massspectrometry. Anal. Chem. 1991, 63 1463.

[16] N. Y. Hsu, W. B Yang, C. H. Wong, Y. C. Lee, R. T. Lee, Y. S. Wang, C. H.Chen. Matrix-assisted laser desorption/ionization mass spectrometryof polysaccharides with 2′,4′,6′-trihydroxyacetophenone as matrix.Rapid. Comm. Mass Spectrom. 2007, 21, 2137.

[17] I. B. Beech, J. Sunner. Biocorrosion: towards understanding interactionsbetween biofilms and metals. Cur. Opinion Biotech. 2004, 15, 181-186.

[18] L. Chi, G. Yue, G. Liu, T. Zhang. Bioproducts fromAureobasidiumpullulans, a biotechnologically important yeast. Appl. Microbiol.Biotechnol. 2009, 82, 793.

[19] R. B. Srivastava, M. Awasthi, M. C. Uprethi, G. N. Mathur. Studies onAureobasidium pullulans forming biofilm on high strengthaluminium alloys, a structural component, in aircraft fuel tanks. Ind.J. Engineer. Mat. Sci. 2006, 13, 135.

[20] H. O. Bouveng, H. Kiessling, B. Lindlberg, J. McKay. Polysaccharideselaborated by Pullularia pullulans. The partial acid hydrolysis of theneutral glucan synthesized from sucrose solutions. Acta Chem.Scand. 1963, 17, 797.

[21] B. McNeil, B. Kristiansen. Temperature effects on polysaccharideformation byAureobasidium pullulans in stirred tanks. EnzymeMicrobiol. Tech. 1990, 12, 521.


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