APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Jan. 1981, p. 268-273 Vol. 41, No. 10099-2240/81/010268-06$02.00/0

Disruption of the Primary Fouling Sequence on Fiber Glass-Reinforced Plastic Submerged in the Marine Environment

DAVID A. CARONt AND JOHN M. SIEBURTH*

Graduate School of Oceantography, University ofRhode Island, Kingston, Rhode Island 02881

Fiber glass-reinforced plastic immersed in an experimental estuarine mesocosmfouled at estimated rates of 0.5, 5.5, and 18.8 ng (wet weight) mm-2 day-' overdays 0 to 2, 2 to 6, and 6 to 14, respectively. Protists, dominated by diatoms,which developed between days 3 and 6 and covered 90% of the undisturbedsurface in 2 weeks, were effectively removed by twice-weekly brushing of thesurface to maintain an immature 3-day bacterial film which covered 12% or lessof the surface and had a biomass 3 orders of magnitude smaller than surfaceswith 2 weeks' unrestricted fouling. Direct brushing of the fiber glass-reinforcedplastic tank walls of experimental estuarine mesocosms minimized the "walleffect" by keeping a surface that maintained a low biomass of a slowly accumu-lating bacterial film rather than a surface which supported the more rapidaccumulation of protists which in turn may induce the settlement of invertebratesand macrophytes.

Microbial fouling of submerged surfaces in thesea (2, 3, 14, 20) is a precursor to macrofoulingin the sea (5, 6, 21). This biological fouling is amajor problem, causing a loss of heat transfer inheat exchangers such as those proposed forocean thermal energy conversion, the loss ofprecision from submerged sensors, an increasein the hull drag of ships, and an undesirableeffect on contained experimental planktonic eco-systems. It is primarily with the latter problemthat this study was concerned, but the resultsare applicable to the limiting of fouling in gen-eral.The use of large-volume enclosures for the

study of biological and chemical processes takingplace in the marine environment has receivedimpetus in recent years (1, 16, 17). The study ofany aquatic system in this manner has intrinsicin it the problem of containment. The introduc-tion of walls around the study system introducesinfluences mediated by the presence of thosewalls, and this "wall effect" is of utmost concernif results obtained in these enclosures are in-tended to be applicable to the natural environ-ment (18, 22).

This study evaluated the feasibility of limitingthe accumulation ofmicroorganisms on the wallsof an experimental estuarine ecosystem by reg-ular brushing. Experiments were carried out atthe facilities of the Marine Ecosystem ResearchLaboratory (MERL) of the University of RhodeIsland (19; M. E. Q. Pilson, C. A. Oviatt, and S.W. Nixon, Proc. Symp. on Microcosms in Eco-

T Present address: Woods Hole Oceanographic Institute,Woods Hole, MA 02543.

logical Research, Augusta, Ga., in press). Theexperimental mesocosms at MERL are cylindri-cal tanks (inner diameter, 1.83 m; water columnheight, 5.0 m) composed of fiber glass-reinforcedplastic. These tanks have been designed to sim-ulate the planktonic ecosystem and its underly-ing benthos in nearby Narragansett Bay as ameans of determining the effect and fate ofvarious pollutants which might enter nearshorewaters.A cleaning procedure was adopted which con-

sists of lowering and raising a circular brushassembly into the cylindrical tank to within 0.5m of the bottom by means of a pulley system(see Fig. 420 in reference 9). The resulting fric-tion between the 5-cm monofilament nylon bris-tles and the tank wall dislodges organisms foul-ing the surface of the wall. This cleaning proce-dure has been conducted at 3- to 4-day intervalsin an attempt to limit the microorganisms colo-nizing the walls and prevent the more obviousfouling by invertebrates and macrophytes. Thepurpose of this paper is to document the effec-tiveness of this brushing procedure in preventingthe development of the fouling community onthe walls of the ecosystem tanks and to pointout its potential for the disruption of fouling onany smooth surface submerged in the sea.

MATERIALS AND METHODSThis investigation was conducted in the late sum-

mer and early autumn months of 1976 when foulingwas maximal. The fouling arrays were made of thesame fiber glass material composing the tank wall, theinnermost surface of which is a cross-linked polysty-rene resin (Beetle Plastics Co., Fall River, Mass.).Disks 10 mm in diameter were cut from fiber glass

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DISRUPTION OF MICROBIAL FOULING 269

sheets and attached to rectangular sheets of fiber glass(20 by 20 cm). Two of these experimental foulingarrays were hung in each of the three control tanks(no. 3, 5, and 7) of the MERL facility. The foulingsurface was oriented vertically facing the center of thetank at a depth of 1.5 m. The experimental tanks usedfor this study were not stressed with pollutants at anytime during this investigation. Just before each sam-pling, one fouling array in each tank was brushed in acleaning procedure which closely mimicked that usedin the tanks, whereas the other array received nocleaning throughout the study period. Immediatelyupon sampling the disks were fixed in 3% glutaralde-hyde in 0.1 M sodium cacodylate buffer plus 0.25 Msucrose and processed in the usual manner by usingcritical point drying for scanning electron microscopy(20). Counts were made by scanning electron micros-copy of both bacterial and protist cells (autotrophicand heterotrophic single-celled eucaryotes) per unitarea by scanning at a 00 angle at a fixed magnification.Replicate counts were made at randomly selected siteson the fouling disk to insure representative results. Inmost cases two or more disks from each array wereobserved. The results from similarly treated foulingdisks held in different experimental tanks showed nosignificant differences and were thus combined in thedetermination of all parameters (i.e., cell density, spe-cies diversity, biomass, and percent cover). Speciesidentification of the protists was based on scanningelectron micrographs of the fouling disks in conjunc-tion with light microscopic examination of organismscolonizing glass microscope slides exposed in the ex-perimental tanks during the study period. Diatoms,the only algal group observed, were identified by ref-erence to Cupp (7) and Hendey (11). Heterotrophiceucaryotes were identified by reference to Grell (10),Kahl (12), and Kudo (13). The surface area of foulingdisks covered by microorganisms and debris was meas-ured for three or more micrographs per fouling disk. Agrid of 100 evenly-spaced points was overlaid on eachmicrograph. The number of these locations coveredby material was used as the estimate of percent cover.Volume estimates were calculated from scanning elec-tron micrographs for each protist species observedbased on its shape and dimensions. For bacterial esti-mates, the cells were grouped into one of four cate-gories depending upon size and shape. A volume wasdetermined for each of these bacterial "types," andthe bacterial volume present per unit area was deter-mined (8, 23). Bacteria occurring in chains or filamentsor both were enumerated separately and placed intothe appropriate category. Biomass for bacterial andprotist populations per unit area, expressed in wetweight, was converted from volume estimates on thebasis of 1 cm3 = 1 g. In instances where protistsstrongly dominated the fouling film, as with the un-cleaned fouling arrays after 10 days of exposure, bac-terial cells were not enumerated because of their lowcontribution to total biomass and because they wereobscured by the larger protists.

RESULTSA simulation of the brushing procedure em-

ployed by MERL was effective in the disruption

of the microbial fouling process on the resinsurface of the fiber glass tank material and re-turned the microbial fouling process to a rela-tively early stage. Figure 1 presents scanningelectron micrographs of the surface when al-lowed to foul naturally (Fig. 1, A, B, and C) andwhen allowed to foul for the same period butbrushed as described just before sampling (Fig.1, D, E, and F). Virtually all protist cells wereremoved by the brushing assembly. Figures 2and 3 show the effect of the brushing process onprotist species diversity and total protist celldensity, respectively. Periodic brushing of thefouling arrays drastically reduced colonization.Unbrushed surface material was densely popu-lated by protists after 16 days of exposure (1,950cells mm-2), whereas material subjected to thebrushing process showed very few (if indeed anydetectable) protists. Day 6 and day 10 values inFig. 3 for brushed arrays fell below the level ofdetectability (1 protist per 2 mm2) by the meth-ods used here. This is further illustrated by thesurface area covered by fouling shown in Fig. 4.Brushing kept the surface cover below 12%,whereas about 90% of the surface was coveredon the unbrushed array. In contrast to the un-brushed fouling arrays, bacterial contribution tothe living biomass on the brushed arrays wassignificant throughout the study period. Thiswas not due to an increase in bacterial biomass(which in fact remained relatively constant), butrather to the more efficient removal of protistsfrom the surface. Figure 5 shows the changes inbiomass per unit area for brushed and un-brushed fouling arrays. The log scale is decep-tive, and although the initial rate of biomassaccumulation appears most rapid and to slowlyfall off to a plateau after day 14, the converse istrue. Total fouling accumulated at a rate of 0.5ng (wet weight) mm-2 day-' over the first 2 days,5.5 ng mm-2 over the next 4 days, and at 18.8 ngmm-2 day-' over the next 8 days.

DISCUSSIONColonization ofthe unbrushed fiber glass-rein-

forced plastic surface during this period (Fig. 1)was strongly dominated by nitzschioid diatomsconsisting of larger Nitzschia-like forms and asmaller fusiform species similar to Cylindro-theca which accounted for the low species diver-sity values of Fig. 2. Pennate diatoms were themajor component of the primary fouling filmthroughout this study, due undoubtedly to thelight intensity at the shallow water depth usedto immerse the test panels and the white colorof the fiber glass wall material. The brushingprocess used at the MERL facility was not onlyeffective in the removal of these gliding pennate

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270 CARON AND SIEBURTH

FIG. 1. Transition of an unrestricted fouling film (A, B, and C) from procaryote to protist domination(fusiform nitzschioid diatoms) occurred between days 3 (A) and 6 (B), whereas larger Nitzschia-like formsappeared by day 16 (C). Twice-weekly brushing of the fiber glass-reinforced plastic substrata (D, E, and F)removed the protists and restricted bacterial colonization to microcrevices where concentrations at days 6 (E)and 16 (F) were similar to those at day 3 (D). Bars, 50 p,m.

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DISRUPTION OF MICROBIAL FOULING 271

10

I-

zw

lf)wita.:

U)

w

C-)w

LLCo

8

6

4

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FIG. 2. Effect ofmechanical disruption on the spe-cies diversity ofprotists attached to fiber glass-rein-forced plastic fouling arrays submerged in an estua-rine mesocosm.

2000

cm 1800E

Effi 1600U/)-J

J 1400w

1200cn

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FIG. 3. Effect ofmechanical disruption on thepop-ulation density ofprotist assemblages, dominated bypennate diatoms, accumulating on fiber glass-rein-forced plastic fouling arrays submerged in an estua-rine mesocosm. Error bars indicate + 1 standarddeviation.

diatoms but also removed many forms with a

high affinity for the substratum. Amoebae, ses-

sile choanoflagellates, suctorians, and vorticellidciliates (21) were also removed by the brushingprocess, as were large amounts of detritus whichaccumulated along with the living microorga-nisms. Stalked bacteria and other bacterialforms possessing attachment organelles such as

fimbrae or pilli likewise showed no resistance tomechanical removal.The fouling process is initiated by bacteria

free in the water and adsorbed onto particulates(2, 4) with the subsequent appearance of protistsand macroorganisms (20, 21). The smooth na-

ture of the resin surface is undoubtedly a largefactor in the high efficiency of removal by thebrushing procedure. The microorganisms are af-forded little protection from the abrasive actionof the brush due to the lack of deep furrows or

fissures. The surface is not completely smoothat the microscopic level, however, and the shal-low microcrevices are the usual location of sed-imented debris and microorganisms which re-

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DAYS OF EXPOSUREFIG. 4. Estimates of percent surface cover for

brushed and unbrushed fiber glass-reinforced plasticfouling arrays submerged in an estuarine mesocosm.

Error bars indicate + I standard deviation.

* UNBRUSHED FOULINGARRAYS

0 BRUSHED FOULINGARRAYS

-, ___a-

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272 CARON AND SIEBURTH

i6

cm

E l

Ecn

20 10-)0u

0 4 8 12 16DAYS OF EXPOSURE

FIG. 5. Effect of mechanical disruption on the liv-ing biomass of the fouling film accumulating on fiberglass-reinforced plastic fouling arrays submerged inan estuarine mesocosm. Biomass is expressed as wetweight; error bars indicate + I standard deviation.

main after the brushing procedure (Fig. 1, D, E,and F).Although the study period was relatively

short, colonization of microorganisms on the un-

brushed fouling disks appeared to have reacheda maximum by day 16. Biomass at this time wasapproximately 150 ng mm-2. Protist species di-versity also appeared to be maximal by day 16(Fig. 2) and may be due to the sloughing of partof the film from the surface at this time. Thishypothesis is also supported by the percentcover values (Fig. 4) for these disks which showa maximum at day 14 for unrestricted (un-brushed) colonization and a drop at day 16.There is also a shift in species dominance at thistime.The gross effect of the brushing procedure

employed at MERL is to effectively remove thefaster-accumulating protists which are second-ary colonizers and leave a primary fouling filmof more slowly accumulating bacteria equal to a

"day 3" stage of colonization. It is possible thatthis mechanical disruption of the microbial foul-ing process may also substantially reduce theattachment of larval invertebrates, since thisprocess has been suggested as an important fac-tor in the conditioning of the substratum forinvertebrate settlement (2, 3, 5, 6, 15, 24). Fur-thermore, the removal of the protists at 3-day

intervals prevented a major film buildup. Usingthe apparent rate of biomass accumulationshown in Fig. 5, we calculated that by day 15unrestricted fouling produced 150 ng (wetweight) mm-2 of wall. Protist removal every 3days maintained a fouling rate of 5.5 ng mm-2day-' (rate calculated for days 2 to 6) throughday 15, with protists contributing 66 ng to theplankton for each mm2 of wall by day 15. Thisis less than half of the contribution of protists inthe unrestricted film. Also, these estimates donot take into account additional contributions ofunrestricted fouling due to sloughing of flhmmaterial into the water and inducement of larvalsettlement. The restricted bacterial film onbrushed surfaces would have a minimal walleffect, affecting only heterotrophic utilization oforganic matter, whereas the mature protist filmdominated by pennate diatoms would not onlycontribute to primary productivity but alsowould falsely influence the measurements usedto determine the activity of planktonic photo-trophs.

Minimization of the biomass of attached mi-croorganisms is essential to the validity of re-search obtained from contained ecosystem proj-ects such as MERL. Unlike controlled ecosystempollution experiments (CEPEX) (16) and otherbag ecosystems anchored in the water column,the placement of MERL tanks on shore not onlyallows greater safety and sampling flexibility butpermits antifouling measures which can con-tinue over extended periods.Mechanical removal of attached microorga-

nisms does have the complication of causing asmall biomass input to the contained watermass. However, fouling organisms affect the en-closed system whether or not they are attached.Periodic removal of the fouling film minimizesthe effect of wall growth to processes within theexperimental system by maintaining a slow foul-ing rate characteristic of the initial fouling se-quence.Mechanical disruption of the fouling film at

regular intervals before extensive fouling takesplace is a novel idea and merits further investi-gation. It should be directly applicable to anysmooth surface such as the fiber glass-reinforcedplastic hulls of pleasure and working craft, thesensing surfaces of submerged instruments, andthe heat-exchanging surfaces of ocean thermalenergy conversion systems.

ACKNOWLEDGMENTS

We thank the members of MERL for their cooperation,especially Diego Alonso for fabricating the brush assembly,and Michael E. Q. Pilson for critiquing the manuscript. PaulE. Hargraves is gratefully acknowledged for his aid in diatomidentification.

* UNBRUSHED FOULING ARRAYSA o BRUSHED FOULING ARRAYS, totalI t & BRUSHED FOULING ARRAYS,

tI bacteria only

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VOL. 41, 1981

The work was supported by National Science Foundationgrant OCE-7401537 and Environmental Protection Agencygrants R80421701 and R803902.

LITERATURE CITED

1. Balch, N., C. M. Boyd, and M. Mullin. 1978. Large-scaletower tank systems. Rapp. P. V. Reun. Cons. Int. Ex-plor. Mer 173:13-21.

2. Characklis, W. G. 1973. Attached microbial growths. I.Attachment and growth. Water Res. 7:1113-1127.

3. Corpe, W. A. 1972. Microfouling: the role of primary filmforming marine bacteria, p. 598-609. In R. F. Acker, B.F. Brown, J. R. DePalma, and W. P. Iverson (ed.),Proceedings of the Third International Congress on

Marine Corrosion and Fouling. National Bureau ofStandards, Gaithersburg, Md.

4. Corpe, W. A. 1974. Periphytic marine bacteria and theformation of microbial films on solid surfaces, p. 397-417. In R. R. Colwell and R. Y. Morita (ed.), Effect ofthe ocean environment on microbial activities. Univer-sity Park Press, Baltimore.

5. Crisp, D. J. 1974. Factors influencing the settlement ofmarine invertebrate larvae, p. 177-265. In P. T. Grantand A. M. Mackie (ed.), Chemoreception in marineorganisms. Academic Press, Inc., London.

6. Crisp, D. J., and J. S. Ryland. 1960. Influence of filmingand of surface texture on the settlement of marineorganisms. Nature (London) 185:119.

7. Cupp, E. E. 1943. Marine plankton diatoms of the westcoast of North America. Bull. Scripps Inst. Oceanogr.5:1-238.

8. Dexter, S. C., J. D. Sullivan, Jr., J. Williams III, andS. W. Watson. 1975. Influence of substrate wettabilityon the attachment of marine bacteria to various sur-

faces. Appl. Microbiol. 30:298-308.9. Environmental Research Laboratory. 1978. MERL

Report, p. 85. Environmental Protection Agency, Nar-ragansett, R.I.

10. Grell, K. G. 1973. Protozoology. Springer-Verlag, NewYork.

11. Hendey, N. I. 1964. An introductory account of thesmaller algae of British waters. Part V: bacillariophy-ceae (diatoms). Fishery investigation series IV. HerMajesty's Stationery Office, London.

DISRUPTION OF MICROBIAL FOULING 273

12. Kahl, A. 1930, 1931, 1932, 1935. Urtiere oder Protozoa. I.Wimpertiere oder Ciliata (Infusoria). In F. Dahl (ed.),Die Tierwelt Deutschlands und der angrenzendenMeersteils. Teils 18, 21, 25, 30. Verlag Fischer, Jena.

13. Kudo, R. R. 1966. Protozoology. Charles C Thomas, Pub-lisher, Springfield, Ill.

14. Marshall, K. C. 1976. Interfaces in microbial ecology.Harvard University Press, Cambridge, Mass.

15. Meadows, P. S., and G. B. Williams. 1963. Settlementof Spirorbis borealis Daudin larvae on surfaces bearingfilms of micro-organisms. Nature (London) 198:610-611.

16. Menzel, D. W., and J. Case. 1977. Concept and design:controlled ecosystem pollution experiment. Bull. Mar.Sci. 27:1-7.

17. Menzel, D. W., and J. H. Steele. 1978. The applicationof plastic enclosures to the study of pelagic marinebiota. Rapp. P. V. Reun. Cons. Int. Explor. Mer 173:7-12.

18. Oviatt, C. A., K. T. Perez, and S. W. Nixon. 1977.Multivariate analysis of experimental marine ecosys-tems. Helgol. Wiss. Meeresunters. 30:30-46.

19. Pilson, M. E. Q., G. A. Vargo, P. Gearing, and J.Gearing. 1977. The Marine Ecosystems Research Lab-oratory: a facility for the investigation of effects andfates of pollutants, p. 513-516. In Proceedings of the2nd National Conference of the Interagency Energy/Environment Research and Development Program,U.S. Environmental Protection Agency, Washington,D.C.

20. Sieburth, J. M. 1975. Microbial seascapes. UniversityPark Press, Baltimore.

21. Sieburth J. M. 1979. Sea Microbes. Oxford UniversityPress, New York.

22. von Bodungen, B., K. von Brockel, V. Smetacek, andB. Zeitzchel. 1976. The plankton tower. I. A structureto study water/sediment interactions in enclosed watercolumns. Mar. Biol. 34:369-372.

23. Watson, S. W., T. J. Novitsky, H. L. Quinby, and F.W. Valois. 1977. Determination of bacterial numberand biomass in the marine environment. Appl. Environ.Microbiol. 33:940-954.

24. Wilson, D. P. 1955. The role of micro-organisms in thesettlement of Ophelia bicornis Savigny. J. Mar. Biol.Assoc. U.K. 34:531-543.

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