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Characterization of biofilm development on artificial substratum in natural water
War. Res. Vol. 27, No. 3, pp. 361-367,1993 Printedin Great Britain.All rightsreserved
0043-1354/93 $6.00+ 0.00 Copyright © 1993PergamonPress Ltd
CHARACTERIZATION OF BIOFILM DEVELOPMENT ON ARTIFICIAL SUBSTRATUM IN NATURAL WATER D. LIU, Y. L. LAU, Y. K. CHAU and G. J. PACEPAVICIUS Rivers Research Branch, National Water Research Institute, 867 Lakeshore Road, P.O. Box 5050, Burlington, Ontario, Canada L7R 4A6 (First received March 1992; accepted in revised form August 1992)
Abstract--In aquatic systems with large specific areas (bottom surface area/volume of water), such as shallow streams or rivers, the growth of biofilms often determines the rate at which environmental contaminants are removed and degraded. Thus, understanding biofilm growth is critical in predicting the ultimate fate of chemicals in the aquatic environment. Through sequential acid hydrolysis and HPLC analysis of biofilms,bacteria and algae, our study provided the first direct biochemical evidence that the origin and mass of a biofilm was mainly derived from bacterial activity, thus justifying the approach of using the major carbohydrate component of microorganisms to approximate the mass of a biofilm. Our data also demonstrated the potential applicability of the carbohydrate method for biofilm mass estimation in various aquatic environments, as biofilm mass can be readily determined in the presence of seawater and hard water. Key words--biofilm development, bacterial biofdm, biomass estimation, carbohydrate
INTRODUCTION Bacterial adhesion to solid submerged objects and subsequent growth often lead to the formation of biofilms. The process of biofilm development is of major importance in aquatic environments. It provides microorganisms living in nutrient-deficient systems, such as oligotrophic waters, with sites where growth and survival can take place (Costerton et al., 1978). Microbial activity resulting from such attached biofilm is also the driving force for many important biogeochemical processes, such as nutrient recycling and degradation of contaminants (Gantzer et al., 1988a). From the perspective of microbial ecology, three distinct types of microbial population exist in the aquatic environment--the dispersed microorganisms (free-living), the fixed bacterial biofilm and the microorganisms attached to other larger organisms such as algae and fish. In general, bacterial biofilm tends to predominate in aquatic systems with large specific areas (bottom surface area/volume of water), such as shallow streams or rivers (Gantzer et al., 1988b). Interestingly, recent studies have unambiguously established that the growth of biofilm often determines the rate at which environmental contaminants are removed and degraded in such systems (Carey et al., 1984; Gantzer et al., 1988a). Thus, understanding the growth of biofilm in natural waters is crucial in predicting the ultimate fate of chemicals in the aquatic environment. As a complex of functional consortia made up of microorganisms, organic and inorganic solids (McFeters, 1984), biofilm is not a single entity.
Consequently, biofilm mass is normally estimated by approximations. These may include a measurement of the biofilm's thickness (La Motta, 1976; Nielsen, 1987), determination of its wet and dry weights (Pedersen, 1982a), and assay for its ATP, protein, polysaccharide and C/N contents (Aftring and Taylor, 1979). In a previous communication (Liu et al., 1992), we described the development of a rapid method for the quantitative estimation of biofilm mass based on the direct measurement of carbohydrate content in biofilms which have been developed on 15 x 25 mm mini glass slides. In the present report, we attempt to characterize the process of biofilm development on artificial substratum in water of natural origin and to provide direct biochemical evidence on the origin of such biofilms. The applicability of the carbohydrate method in quantifying biofilm mass in other related areas is also discussed. MATERIALSAND METHODS Cultivation of cultures Bacillus cereus and mixed bacterial culture (from Hamilton Harbour water) were all grown on an enrichment medium with the following ingredients (g/l): 1 g each of nutrient broth, peptone and yeast extract; 2 g each of glucose and sodium acetate; K2HPO4, 2.64 g; KH2HPO(, 1.64 g; distilled water, 1 litre. The medium was sterilized by autoclaving at 121°C for 15 min. After a 20-h growth on a rotary shaker (220 rpm) at room temperature (21°C), the bacterial cells were harvested by centrifugation (10,000g for 15 min), washed once in cold 0.02 M phosphate buffer (pH 7.0) and were used directly in the acid hydrolysis for their carbohydrate components. The unicellular green alga Selenastrum capricornutum (ATCC 22662) was grown according to the procedure of Blaise (1986). The algal cells were also harvested by centrifugation (3000g for 15 min) 361
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and washed once in cold phosphate buffer prior to acid hydrolysis for carbohydrate components. It would have been ideal to grow the bacterial cultures on glass slides instead of on enriched media. However, acid hydrolysis of bacterial biomass for their mono sugar components requires a large amount of sample, usually 1 g or more. To obtain such quantities of samples, the bacteria had to be grown in very high concentration nutrient medium. Unfortunately, under such conditions, the bacteria would only grow in a dispersed state (i.e. would not form biofilms).
Seawater and hard water Synthetic seawater (Jones et al., 1976) had the following ingredients (g/l): NaCI, 23.476; MgCI:. 6H20, 10.635; Na2SO4, 3.917; CaC12, 1.102; KCI, 0.664; NaHCO3, 0.192; KBr, 0.096; HH3BO3,0.026; SrCI2 • 6I-I20, 0.040; NaF; 0.003. The artificial hard water contained the following ingredients (g/l): CAC12.2.51-I20, 0.20; MgSO4'6H20, 0.20; Na2CO3, 0.20; NaHCO3, 0.10.
Preparation of mini slides Mini glass slides (15 x 25 x 0.95 mm) used for supporting the development of biofilm were prepared from precleaned standard microscope slides (25 x 75 x 0.95 mm) using a sharp diamond pencil. After soaking overnight in 5% detergent solution of Contrad 70 (Canlab, Toronto), rinsing and oven-drying, the mini slides were ready for use in the biofilm experiments. To detect the growth of biofilm in the aquatic environment, the slides were laid flat on the bottom of an outdoor flume or a laboratory tank/reactor to simulate the biofilm growth in a stream bed. Microscopic examination revealed that biofilm development was found only on the top side of the slides, i.e. on the side in contact with the water.
Laboratory studies on biofilm growth Laboratory studies on biofilm growth were conducted using a large size tank (200 x 250 x 100 cm) which has a holding capacity of approx. 30001. To more closely simulate the growth of biofilm in a natural aquatic system, water from Hamilton Harbour was used directly in the experiments without any nutrient addition. Fresh harbour water was fed continuously at a rate of approx. 2001. per day to the tank via a pumping device. At predetermined time intervals, several mini slides were removed from the tank for microscopic examination and for analysis of total carbohydrate content. Since the laboratory was poorly illuminated, no algal growth was expected and none was observed in any of the samples.
Flume study of biofilm growth In an attempt to simulate natural fiver bed conditions, biofilms were grown in an outdoor open channel 0.2 m wide and 2 m long, with 0.2 m high sidewalls. Water was pumped from Hamilton Harbour into a headtank which fed into the open channel. The depth of flow in the channel was controlled by a vertically sliding tail gate installed at the downstream end. A 3 x 3 x 1 m tailbox was used to catch the water exiting the channel and to guide it back into the harbour via a 90° V-notch weir. The bottom of the channel was covered with glass. In sections, the glass cover consisted of 15 x 25mm mini slides and 25 x 75mm microscope slides, both of which could easily be removed for analysis. To start an experiment, water was pumped into the headtank at the desired flow rate and the tailgate was adjusted to give the desired flow depth. The mini slides and microscope slides were then placed into the channel bottom to begin the growth of biofilms. With the exception of the artificial substratum, this setup provided conditions for biofilm growth closely resembling those in natural rivers.
Assay of carbohydrate content in biofilm The content of total carbohydrate in the biofilm developed on the mini slides was determined by a recently
developed biochemical method (Liu et al., 1992). To perform the test, the mini slide was allowed to react with the phenol--sulphuric acid reagent in a 20 x 150 mm test tube and the absorbance spectrum of the reaction mixture was scanned in a spectrophotometer from 650 to 350 nm for the absorbance maximum against a reagent blank. As the development of biofilm may not be homogeneous, a minimum of 3 mini slides was used to obtain the average in each assay.
Hydrolysis and analysis of biofilm, bacteria and algae The hydrolysis and analysis of biofilm, bacteria and algae for their carbohydrate components followed conventional procedures (Borchardt and Piper, 1970; Binder, 1980). Approximately I g of sample (wet weight) was placed into a 250-ml Erlenmeyer flask containing 3 ml of 72% HH2SO4. The mixture was incubated on a rotary shaker (150 rpm) for I h at room temperature (21°C) for primary hydrolysis of polysaccharides (i.e. formation of oligosaccharides). The resulting hydrolysate was then diluted with 84 ml of distilled water and was autoclaved for I h at 121°C to yield the monomeric sugars (secondary hydrolysis). Upon cooling, the resulting hydrolysate was neutralized with a saturated solution of barium hydroxide [70 g of Ba(OH)2.8HH20 per litre] to pH 5.5 under vigorous agitation. The neutralized hydrolysates were then centrifuged at 8000g for 10 min to remove the formed barium sulphate and the supernatant was further concentrated down to approx. 10 ml on a rotary evaporator at 60°C. The concentrate was centrifuged again at 3000 g to remove any remaining barium sulphate and 25 #1 of the supernatant was directly analysed for sugars on a high performance liquid chromatograph (Waters Model 600E) equipped with a photodiode array detector (Water Model 900). The 3.9 x 300 mm Waters carbohydrate column, packed with silica gel with an amine functionality, was operated at ambient temperature (21°C). The mobile phase was acetonitrile: water (80:20) with the flow rate at 2 ml/min. Since carbohydrates absorb only at a wavelength lower than 200 rim, the photodiode array detector was set at 190 nm (Binder, 1980).
Microscopic examination of biofilm At predetermined time intervals, microscope slides were taken to the laboratory for microscopic examination of biofilm growth and development. A Leitz (model Dialux) phase contrast microscope equipped with an automatic micrograph attachment was used to assess and record the biofilm development on test microscope slides. The sequential development of biofilm on the test slides was photographed using a high speed (ASA 3200) Polaroid type 107 film. RESULTS T h e direct c a r b o h y d r a t e m e t h o d (Liu et al., 1992) was f o u n d to be highly sensitive a n d c o n v e n i e n t w h e n used in assessing the q u a n t i t a t i v e g r o w t h o f biofilm in n a t u r a l water. After l - h exposure to the flowing H a m i l t o n H a r b o u r w a t e r (3.4 cm/s) in the o u t d o o r o p e n channel, biofilm g r o w t h (in terms o f total c a r b o h y d r a t e ) o n test mini slides was detected by the m e t h o d . This p r o m p t e d us to examine the slides u n d e r a phase c o n t r a s t microscope a n d the observations revealed the occurrence o f biofilm g r o w t h o n the slides. Because o f its ability to provide rapid qualitative i n f o r m a t i o n a b o u t biofilm development, microscopic e x a m i n a t i o n was, therefore, routinely u n d e r t a k e n prior to each c a r b o h y d r a t e d e t e r m i n a t i o n for biofilm estimation.
Characterization of biofilm development Biofilm growth in natural water was studied both indoors and outdoors in an attempt to delineate the nature of biofilm development. Apparently the onset and the subsequent development of a natural biofilm is an intricate process which is sensitive to even some minor or subtle changes in the experimental condition. For example, the same Hamilton Harbour water was used in all the experiments and yet biofilms developed indoors appeared to differ significantly from the ones grown outdoors. The biofilm developed indoors always had a slower growth rate and consisted of organisms of simpler trophic level, with mainly bacteria as the predominant microorganisms (Fig. 1). On the other hand, biofilm developed in the outdoor open channel system had a much faster growth rate and was characterized with a wider spectrum of organisms. Besides bacteria, many different kinds of other organisms such as protozoa, diatoms, rotifers, algae and worms were also present (Fig. 2). It was observed that bacteria were predominant at the beginning of biofilm development in both the indoor tank and the outdoor flume. In the early stages, a biofilm from the outdoor flume would closely resemble that shown in Fig. 1 from the indoor tank. However, whereas bacteria remained dominant in the indoor system throughout the entire period of growth, the structure of the biofilms in the outdoor flume became much more complex as growth progressed,
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Fig. 2. Phase contrast micrographs (500 x magnification) of biofilm after 26 h exposure to the flowing (3.4 cm/s) Hamilton Harbour water in the outdoor open channel on 4 July 199I, (A) upper layer and (B) lower layer.
with many other organisms growing on top of the existing first layer of cells (Fig. 2). Other studies (Jordan and Staley, 1976; Burkholder and Wetzel, 1989) have also found that bacteria were the primary colonizers of both artificial and natural substratum, to be gradually supplemented by other organisms. To delineate the origin of natural biofilms, acid hydrolysis was conducted on samples of biofilm, bacterial and algal cells for the composition of their polysaccharides [Fig. 3(A)]. The appearance of discrete peaks in the secondary acid hydrolysates indicates the successful hydrolysis of these polysaccharides into the simpler monomeric sugars [Fig. 3(B)]. This deduction was further confirmed by HPLC analysis of these hydrolysates for monomeric sugars. Figure 4 shows HPLC profiles of the secondary acid hydrolysates prepared from natural biofilm, mixed bacterial cultures, pure bacterial culture (Bacillus cereus) and pure algal culture (Selenastrum capricornutum). Based on the comparison of retention times and peak-profile matching, it can be tentatively concluded that the origin of natural biofilm (outdoor open channel) was mainly developed from the bacterial activity. Both the pure and mixed bacterial preparations were found to match Fig. 1. Phase contrast micrographs (500 × magnification) of well with the biofilm hydrolysate. The algal hybiofilm after: (A) 5 h and (B) 23 h, exposure to Hamilton drolysate could match with only one peak of the biofilm's hydrolysate, thus implying a minor role Harbour water in an indoor tank on 22 April 1991. WR 27/3~C
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Fig. 3. Absorbance spectra of carbohydrates (phenol-sulphuric acid) from natural biofilm, B. cereus and S. capricornumm, (A) before acid hydrolysis and (B) after secondary acid hydrolysis. of the algal population in the development of a natural biofilm. However, we must emphasize that Bacillus cereus and Selenastrum capricornutum are not the predominant species found in natural biofilms and therefore the present observations may not necessarily apply. We have used these cultures in our experiments because of their ease of growth and their availability, even though they are not the ideal choices. Marine environment is frequently the final sink for many toxic substances and thus experiments were conducted to study the feasibility of using the direct carbohydrate method for the estimation of biofilm mass in the presence of seawater and hard water. The biofilm was grown in the outdoor flume and the analysis for carbohydrate was carried out in the presence of synthetic sea water and hard water. Figure 5 shows that the method can be successfully conducted in the presence of seawater or hard water. The high concentration of inorganics from seawater (10% full strength and higher) may sometimes cause the formation of fine suspended particles in the carbohydrate reaction mixture, but this interference
can be easily removed by a low-speed centrifugation (1000 rpm for 5 min) prior to the measurement for carbohydrate. DISCUSSION This study and literature data (Pedcrsen, 1982b) have highlighted some of the problems involved in the study of biofilm growth in natural water. Some of the factors that control the initiation and the subsequent biofdm development need to be resolved. At the present time, it seems impossible to duplicate the development process of a natural biofilm in the setting of an indoor laboratory. For example, the same Hamilton Harbour water when used in the indoor laboratory tank would produce biofilms vastly different from those developed in the outdoor open channel system. Very likely the nature of biofilms that develop in an aquatic environment may depend on many factors, especially the characteristics of that particular aquatic system and the water itself (Pedersen, 1982a). For example, certain freshwater bacteria have been shown to have a preference for
Characterization of biofilm development
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attachment to hydrophobic surfaces such as Teflon and PVC (Pringie and Fletcher, 1983). Consequently, biofilms grown on natural substratum may not necessarily have the same development as the ones grown on glass surfaces in our study. Our study results (Fig. 1) as well as the observations of Dexter et al. (1975) indicate that biofilm development at the early stage was non-homogenous. This is different from some earlier observations such
365
as those of Pedersen (1982b). This difference is likely due to the fact that biofilms become more uniform in texture and appearance as they mature. As the biofilm developed from initial patches of bacteria, more cells and exopolymers (polysaccharides) would accumulate and fill the void space. In the mature biofilm, several distinct layers of cells made up of different organisms could be seen in the biofilm matrix (Fig. 2). Consequently, a mature biofilm would take on the macroscopic appearance of a gel which contributes to its appearance of uniformity in texture. A similar observation on the appearance of mature biofilm was also reported by McFeters (1984). The high content of polysaccharides in a mature biofilm is also likely to be responsible for a biofilm's gelling or water-holding properties. Certain polysaccharides have been known to be capable of forming aqueous gels at very low concentrations (Aftring and Taylor, 1979). Apparently, this gelatinous matrix may also act as a trap for debris and nutrients (Pedersen, 1990), thus contributing to the overall physical, chemical and biological properties of a mature biofilm. From the viewpoint of microbial physiology, the majority of the biofilm studies in the literature were in the category of accelerated biofilm growth which involved the use of high concentrations of organic nutrients to promote rapid growth. The organic nutrients employed included glucose (La Motta, 1976; Characklis et al., 1982; Gantzer et al., 1988b), glycerol (Gilbert et al., 1989), acetate/citrate (Nielsen, 1987), giucose/peptone (Kuroda et al., 1988) and sucrose/yeast extract (McCoy et al., 1981). These chemostat approaches have provided some useful information on the kinetics of biofilm growth, notably in the design of wastewater treatment facilities. However, the utilization of organic nutrient concentrations that are too high to be realistic for natural biofilms would limit their usefulness for predicting the growth rate of natural biofilm in the real, nutrient-poor, aquatic environment. Perhaps the most valuable contribution of this study to the knowledge of biofilm growth was the elucidation of the origin of the natural biofilm. Many organisms including bacteria, protozoa, fungi and algae have been detected in natural biofilms (McCoy et al., 1981; Pedersen, 1982a, b, 1990). However, their relative importance in the development of a natural biofilm was inconclusive. In this study we provided the first direct biochemical evidence that the origin of natural biofilm was mainly derived from bacterial activity, i.e. bacteria being the major component of biofilms which were established on glass slides with water of natural origin. Based on microscopic observations, Pedersen (1982a) also concluded that algae were not involved in biofilm development in the marine environment. Biofilms contain relatively large quantities of polysaccharides (Characklis et al., 1982) and methods based on the reaction of carbohydrate have been
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3,4-dichlorophenol in a fluvial system. Can. J. Physiol. Pharmac. 62, 971-975. Characklis W. G., Trulear M. G., Bryers J. D. and ZeiverN. (1982) Dynamics of biofilm processes: methods. War. Res. 16, 1207-1216. Costerton J. W., Gessey G. G. and Cheng K. J. (1978) How bacteria stick. Scient. Am. 238, 86-95. Dexter S. C., Sullivan J. D. Jr, Williams J. III and Watson S. W. (1975) Influence of substrate wettability on the attachment of marine bacteria to various surfaces. Appl. Microbioi. 30, 298-308. Gantzer C. J., Kollig H. P., Rittmann B. E. and Lewis D. L. (1988a) Predicting the rate of trace-organic compound removal by natural biofilm. Wat. Res. 22, 191-200. Gantzer C. J., Rittmann B. E. and Herrick E. E. (1988b) Mass transport to streambed biofilm. Wat. Res. 22, 709-722. Gilbert P., Allison D. G., Evans D. J., Handley P. S. and Brown M. R. W. (1989) Growth rate control of adherent bacterial populations. Appl. envir. Microbiol. 55, 1308-1311. Jones G. E., Royle L. G. and Murray L. (1976) The assimilation of trace metal ions by the marine bacteria, Arthrobacter marinus and Pseudomonas cuprodurans. In P,EFERENCT~ 0 Proceedings of the Third International Biodegradation Symposium (Edited by Sharpley J. M. and Kaplan A. M.), Aftrin8 R. P. and Taylor B. F. (1979) Assessment of pp. 889-898. Applied Science, London. microbial fouling in an ocean thermal energy conversion Jordan T. L. and Staley J. T. (1976) Electron microscopy experiment. Appl. envir. Microbiol. 311, 734-739. of succession in the periphyton community of Lake Binder H. (1980) Separation of monosaccharides by highWashington. Microb. Ecol. 2, 241-251. performance liquid chromatography: comparison of Kuroda M., Yuzawa M., Sakakibara Y. and Okamura M. ultraviolet and refractive index detection. J. Chromatogr. (1988) Methanogenic bacteria adhered to solid supports. 189, 414--420. Wat. Pea. 22, 653-656. Blaise C. R. (1986) Micromethod for acute aquatic toxicity assessment using the green alga Selenastrum capricornu- La Motta E. J. (1976) Kinetics of growth and substrate uptake in a biological film system. Appl. etwir. Microbiol. turn. Toxic. Assess. 1, 377-385. 31, 286--293. Borchardt L. G. and Piper C. V. (1970) A gas chromatographic method for carbohydrates as alditol-acetates. Liu D., Lau Y. L., Clmu Y. K. and Pacepavicius G. J. (1992) A simple technique for the determination of bioTappi $3, 257-260. film formation and growth. NWRI Contribution Burkholder J. M. and Wetzel R. G. (1989) Microbial 92-102, National Water Research Institute, Burlington, colonization on natural and artificial macrophytes in Ontario. a phosphorus-limited hardwater lake. J. Phyeol. 25, McCoy W. F., Bryen J. D., Robbins J. and Costerton J. W. 55--65. (1981) Observation of fouling biofilm formation. Can. J. Carey J. H., Fox M. E., Browniec B. G., Metcalfe J. L. and MicrobioL 27, 910-917. Platford R. F. (1984) Disappearance kinetics of 2,4- and
utilized to quantify biofdm mass (Aftring and Taylor, 1979; Characklis et al., 1982). However the physical removal (scrapping or vacuuming) of biofilm from the supporting substratum is required prior to carbohydrate determination for all these procedures. Difficulties have been experienced in the removal of biofilm from the supporting substratum (Aftring and Taylor, 1979). This difficulty is completely eliminated in our direct procedure for the estimation of biofilm mass, thus greatly enhancing the method's simplicity and sensitivity. Moreover, the direct method is very robust and the test can be readily applied to biofilm samples from the marine environment and alkaline lakes. Biofilms can develop on almost any surface exposed to an aqueous environment. Biofilm growth in natural environments is generally slow and our sensitive method can be useful in the estimation of biofilm mass.
Characterization of biofilm development McFeters (3. A. (1984) Biofilm development and its consequences. In Microbial Adhesion and Aggregation (Edited by Marshall K. C.), pp. 109-124, Springer, Berlin. Nielsen P. H. (1987) Biofilm dynamics and kinetics during high-rate sulfate reduction under anaerobic conditions. Appl. envir. Microbiol. 53, 27-32. Pedersen K. (1982a) Method for studying microbial biofilms in flowing-water systems. Appl. envir. Microbiol. 43, 6--13.
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Pedersen K. (1982b) Factors regulating microbial biofilm development in a system with slowly flowing seawater. Appl. envir. Microbioi. 44, 1196-1204. Pedersen K. (1990) Biofilm development on stainless steel and PVC surfaces in drinking water. War. Res. 24, 239-243. Pringle J. H. and Fletcher M. (1983) Influence of substratum wettability on attachment of freshwater bacteria to solid surfaces. Appl. envir. Microbiol. 45, 811-817.
0043-1354/93 $6.00+ 0.00 Copyright © 1993PergamonPress Ltd
CHARACTERIZATION OF BIOFILM DEVELOPMENT ON ARTIFICIAL SUBSTRATUM IN NATURAL WATER D. LIU, Y. L. LAU, Y. K. CHAU and G. J. PACEPAVICIUS Rivers Research Branch, National Water Research Institute, 867 Lakeshore Road, P.O. Box 5050, Burlington, Ontario, Canada L7R 4A6 (First received March 1992; accepted in revised form August 1992)
Abstract--In aquatic systems with large specific areas (bottom surface area/volume of water), such as shallow streams or rivers, the growth of biofilms often determines the rate at which environmental contaminants are removed and degraded. Thus, understanding biofilm growth is critical in predicting the ultimate fate of chemicals in the aquatic environment. Through sequential acid hydrolysis and HPLC analysis of biofilms,bacteria and algae, our study provided the first direct biochemical evidence that the origin and mass of a biofilm was mainly derived from bacterial activity, thus justifying the approach of using the major carbohydrate component of microorganisms to approximate the mass of a biofilm. Our data also demonstrated the potential applicability of the carbohydrate method for biofilm mass estimation in various aquatic environments, as biofilm mass can be readily determined in the presence of seawater and hard water. Key words--biofilm development, bacterial biofdm, biomass estimation, carbohydrate
INTRODUCTION Bacterial adhesion to solid submerged objects and subsequent growth often lead to the formation of biofilms. The process of biofilm development is of major importance in aquatic environments. It provides microorganisms living in nutrient-deficient systems, such as oligotrophic waters, with sites where growth and survival can take place (Costerton et al., 1978). Microbial activity resulting from such attached biofilm is also the driving force for many important biogeochemical processes, such as nutrient recycling and degradation of contaminants (Gantzer et al., 1988a). From the perspective of microbial ecology, three distinct types of microbial population exist in the aquatic environment--the dispersed microorganisms (free-living), the fixed bacterial biofilm and the microorganisms attached to other larger organisms such as algae and fish. In general, bacterial biofilm tends to predominate in aquatic systems with large specific areas (bottom surface area/volume of water), such as shallow streams or rivers (Gantzer et al., 1988b). Interestingly, recent studies have unambiguously established that the growth of biofilm often determines the rate at which environmental contaminants are removed and degraded in such systems (Carey et al., 1984; Gantzer et al., 1988a). Thus, understanding the growth of biofilm in natural waters is crucial in predicting the ultimate fate of chemicals in the aquatic environment. As a complex of functional consortia made up of microorganisms, organic and inorganic solids (McFeters, 1984), biofilm is not a single entity.
Consequently, biofilm mass is normally estimated by approximations. These may include a measurement of the biofilm's thickness (La Motta, 1976; Nielsen, 1987), determination of its wet and dry weights (Pedersen, 1982a), and assay for its ATP, protein, polysaccharide and C/N contents (Aftring and Taylor, 1979). In a previous communication (Liu et al., 1992), we described the development of a rapid method for the quantitative estimation of biofilm mass based on the direct measurement of carbohydrate content in biofilms which have been developed on 15 x 25 mm mini glass slides. In the present report, we attempt to characterize the process of biofilm development on artificial substratum in water of natural origin and to provide direct biochemical evidence on the origin of such biofilms. The applicability of the carbohydrate method in quantifying biofilm mass in other related areas is also discussed. MATERIALSAND METHODS Cultivation of cultures Bacillus cereus and mixed bacterial culture (from Hamilton Harbour water) were all grown on an enrichment medium with the following ingredients (g/l): 1 g each of nutrient broth, peptone and yeast extract; 2 g each of glucose and sodium acetate; K2HPO4, 2.64 g; KH2HPO(, 1.64 g; distilled water, 1 litre. The medium was sterilized by autoclaving at 121°C for 15 min. After a 20-h growth on a rotary shaker (220 rpm) at room temperature (21°C), the bacterial cells were harvested by centrifugation (10,000g for 15 min), washed once in cold 0.02 M phosphate buffer (pH 7.0) and were used directly in the acid hydrolysis for their carbohydrate components. The unicellular green alga Selenastrum capricornutum (ATCC 22662) was grown according to the procedure of Blaise (1986). The algal cells were also harvested by centrifugation (3000g for 15 min) 361
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and washed once in cold phosphate buffer prior to acid hydrolysis for carbohydrate components. It would have been ideal to grow the bacterial cultures on glass slides instead of on enriched media. However, acid hydrolysis of bacterial biomass for their mono sugar components requires a large amount of sample, usually 1 g or more. To obtain such quantities of samples, the bacteria had to be grown in very high concentration nutrient medium. Unfortunately, under such conditions, the bacteria would only grow in a dispersed state (i.e. would not form biofilms).
Seawater and hard water Synthetic seawater (Jones et al., 1976) had the following ingredients (g/l): NaCI, 23.476; MgCI:. 6H20, 10.635; Na2SO4, 3.917; CaC12, 1.102; KCI, 0.664; NaHCO3, 0.192; KBr, 0.096; HH3BO3,0.026; SrCI2 • 6I-I20, 0.040; NaF; 0.003. The artificial hard water contained the following ingredients (g/l): CAC12.2.51-I20, 0.20; MgSO4'6H20, 0.20; Na2CO3, 0.20; NaHCO3, 0.10.
Preparation of mini slides Mini glass slides (15 x 25 x 0.95 mm) used for supporting the development of biofilm were prepared from precleaned standard microscope slides (25 x 75 x 0.95 mm) using a sharp diamond pencil. After soaking overnight in 5% detergent solution of Contrad 70 (Canlab, Toronto), rinsing and oven-drying, the mini slides were ready for use in the biofilm experiments. To detect the growth of biofilm in the aquatic environment, the slides were laid flat on the bottom of an outdoor flume or a laboratory tank/reactor to simulate the biofilm growth in a stream bed. Microscopic examination revealed that biofilm development was found only on the top side of the slides, i.e. on the side in contact with the water.
Laboratory studies on biofilm growth Laboratory studies on biofilm growth were conducted using a large size tank (200 x 250 x 100 cm) which has a holding capacity of approx. 30001. To more closely simulate the growth of biofilm in a natural aquatic system, water from Hamilton Harbour was used directly in the experiments without any nutrient addition. Fresh harbour water was fed continuously at a rate of approx. 2001. per day to the tank via a pumping device. At predetermined time intervals, several mini slides were removed from the tank for microscopic examination and for analysis of total carbohydrate content. Since the laboratory was poorly illuminated, no algal growth was expected and none was observed in any of the samples.
Flume study of biofilm growth In an attempt to simulate natural fiver bed conditions, biofilms were grown in an outdoor open channel 0.2 m wide and 2 m long, with 0.2 m high sidewalls. Water was pumped from Hamilton Harbour into a headtank which fed into the open channel. The depth of flow in the channel was controlled by a vertically sliding tail gate installed at the downstream end. A 3 x 3 x 1 m tailbox was used to catch the water exiting the channel and to guide it back into the harbour via a 90° V-notch weir. The bottom of the channel was covered with glass. In sections, the glass cover consisted of 15 x 25mm mini slides and 25 x 75mm microscope slides, both of which could easily be removed for analysis. To start an experiment, water was pumped into the headtank at the desired flow rate and the tailgate was adjusted to give the desired flow depth. The mini slides and microscope slides were then placed into the channel bottom to begin the growth of biofilms. With the exception of the artificial substratum, this setup provided conditions for biofilm growth closely resembling those in natural rivers.
Assay of carbohydrate content in biofilm The content of total carbohydrate in the biofilm developed on the mini slides was determined by a recently
developed biochemical method (Liu et al., 1992). To perform the test, the mini slide was allowed to react with the phenol--sulphuric acid reagent in a 20 x 150 mm test tube and the absorbance spectrum of the reaction mixture was scanned in a spectrophotometer from 650 to 350 nm for the absorbance maximum against a reagent blank. As the development of biofilm may not be homogeneous, a minimum of 3 mini slides was used to obtain the average in each assay.
Hydrolysis and analysis of biofilm, bacteria and algae The hydrolysis and analysis of biofilm, bacteria and algae for their carbohydrate components followed conventional procedures (Borchardt and Piper, 1970; Binder, 1980). Approximately I g of sample (wet weight) was placed into a 250-ml Erlenmeyer flask containing 3 ml of 72% HH2SO4. The mixture was incubated on a rotary shaker (150 rpm) for I h at room temperature (21°C) for primary hydrolysis of polysaccharides (i.e. formation of oligosaccharides). The resulting hydrolysate was then diluted with 84 ml of distilled water and was autoclaved for I h at 121°C to yield the monomeric sugars (secondary hydrolysis). Upon cooling, the resulting hydrolysate was neutralized with a saturated solution of barium hydroxide [70 g of Ba(OH)2.8HH20 per litre] to pH 5.5 under vigorous agitation. The neutralized hydrolysates were then centrifuged at 8000g for 10 min to remove the formed barium sulphate and the supernatant was further concentrated down to approx. 10 ml on a rotary evaporator at 60°C. The concentrate was centrifuged again at 3000 g to remove any remaining barium sulphate and 25 #1 of the supernatant was directly analysed for sugars on a high performance liquid chromatograph (Waters Model 600E) equipped with a photodiode array detector (Water Model 900). The 3.9 x 300 mm Waters carbohydrate column, packed with silica gel with an amine functionality, was operated at ambient temperature (21°C). The mobile phase was acetonitrile: water (80:20) with the flow rate at 2 ml/min. Since carbohydrates absorb only at a wavelength lower than 200 rim, the photodiode array detector was set at 190 nm (Binder, 1980).
Microscopic examination of biofilm At predetermined time intervals, microscope slides were taken to the laboratory for microscopic examination of biofilm growth and development. A Leitz (model Dialux) phase contrast microscope equipped with an automatic micrograph attachment was used to assess and record the biofilm development on test microscope slides. The sequential development of biofilm on the test slides was photographed using a high speed (ASA 3200) Polaroid type 107 film. RESULTS T h e direct c a r b o h y d r a t e m e t h o d (Liu et al., 1992) was f o u n d to be highly sensitive a n d c o n v e n i e n t w h e n used in assessing the q u a n t i t a t i v e g r o w t h o f biofilm in n a t u r a l water. After l - h exposure to the flowing H a m i l t o n H a r b o u r w a t e r (3.4 cm/s) in the o u t d o o r o p e n channel, biofilm g r o w t h (in terms o f total c a r b o h y d r a t e ) o n test mini slides was detected by the m e t h o d . This p r o m p t e d us to examine the slides u n d e r a phase c o n t r a s t microscope a n d the observations revealed the occurrence o f biofilm g r o w t h o n the slides. Because o f its ability to provide rapid qualitative i n f o r m a t i o n a b o u t biofilm development, microscopic e x a m i n a t i o n was, therefore, routinely u n d e r t a k e n prior to each c a r b o h y d r a t e d e t e r m i n a t i o n for biofilm estimation.
Characterization of biofilm development Biofilm growth in natural water was studied both indoors and outdoors in an attempt to delineate the nature of biofilm development. Apparently the onset and the subsequent development of a natural biofilm is an intricate process which is sensitive to even some minor or subtle changes in the experimental condition. For example, the same Hamilton Harbour water was used in all the experiments and yet biofilms developed indoors appeared to differ significantly from the ones grown outdoors. The biofilm developed indoors always had a slower growth rate and consisted of organisms of simpler trophic level, with mainly bacteria as the predominant microorganisms (Fig. 1). On the other hand, biofilm developed in the outdoor open channel system had a much faster growth rate and was characterized with a wider spectrum of organisms. Besides bacteria, many different kinds of other organisms such as protozoa, diatoms, rotifers, algae and worms were also present (Fig. 2). It was observed that bacteria were predominant at the beginning of biofilm development in both the indoor tank and the outdoor flume. In the early stages, a biofilm from the outdoor flume would closely resemble that shown in Fig. 1 from the indoor tank. However, whereas bacteria remained dominant in the indoor system throughout the entire period of growth, the structure of the biofilms in the outdoor flume became much more complex as growth progressed,
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Fig. 2. Phase contrast micrographs (500 x magnification) of biofilm after 26 h exposure to the flowing (3.4 cm/s) Hamilton Harbour water in the outdoor open channel on 4 July 199I, (A) upper layer and (B) lower layer.
with many other organisms growing on top of the existing first layer of cells (Fig. 2). Other studies (Jordan and Staley, 1976; Burkholder and Wetzel, 1989) have also found that bacteria were the primary colonizers of both artificial and natural substratum, to be gradually supplemented by other organisms. To delineate the origin of natural biofilms, acid hydrolysis was conducted on samples of biofilm, bacterial and algal cells for the composition of their polysaccharides [Fig. 3(A)]. The appearance of discrete peaks in the secondary acid hydrolysates indicates the successful hydrolysis of these polysaccharides into the simpler monomeric sugars [Fig. 3(B)]. This deduction was further confirmed by HPLC analysis of these hydrolysates for monomeric sugars. Figure 4 shows HPLC profiles of the secondary acid hydrolysates prepared from natural biofilm, mixed bacterial cultures, pure bacterial culture (Bacillus cereus) and pure algal culture (Selenastrum capricornutum). Based on the comparison of retention times and peak-profile matching, it can be tentatively concluded that the origin of natural biofilm (outdoor open channel) was mainly developed from the bacterial activity. Both the pure and mixed bacterial preparations were found to match Fig. 1. Phase contrast micrographs (500 × magnification) of well with the biofilm hydrolysate. The algal hybiofilm after: (A) 5 h and (B) 23 h, exposure to Hamilton drolysate could match with only one peak of the biofilm's hydrolysate, thus implying a minor role Harbour water in an indoor tank on 22 April 1991. WR 27/3~C
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can be easily removed by a low-speed centrifugation (1000 rpm for 5 min) prior to the measurement for carbohydrate. DISCUSSION This study and literature data (Pedcrsen, 1982b) have highlighted some of the problems involved in the study of biofilm growth in natural water. Some of the factors that control the initiation and the subsequent biofdm development need to be resolved. At the present time, it seems impossible to duplicate the development process of a natural biofilm in the setting of an indoor laboratory. For example, the same Hamilton Harbour water when used in the indoor laboratory tank would produce biofilms vastly different from those developed in the outdoor open channel system. Very likely the nature of biofilms that develop in an aquatic environment may depend on many factors, especially the characteristics of that particular aquatic system and the water itself (Pedersen, 1982a). For example, certain freshwater bacteria have been shown to have a preference for
Characterization of biofilm development
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attachment to hydrophobic surfaces such as Teflon and PVC (Pringie and Fletcher, 1983). Consequently, biofilms grown on natural substratum may not necessarily have the same development as the ones grown on glass surfaces in our study. Our study results (Fig. 1) as well as the observations of Dexter et al. (1975) indicate that biofilm development at the early stage was non-homogenous. This is different from some earlier observations such
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as those of Pedersen (1982b). This difference is likely due to the fact that biofilms become more uniform in texture and appearance as they mature. As the biofilm developed from initial patches of bacteria, more cells and exopolymers (polysaccharides) would accumulate and fill the void space. In the mature biofilm, several distinct layers of cells made up of different organisms could be seen in the biofilm matrix (Fig. 2). Consequently, a mature biofilm would take on the macroscopic appearance of a gel which contributes to its appearance of uniformity in texture. A similar observation on the appearance of mature biofilm was also reported by McFeters (1984). The high content of polysaccharides in a mature biofilm is also likely to be responsible for a biofilm's gelling or water-holding properties. Certain polysaccharides have been known to be capable of forming aqueous gels at very low concentrations (Aftring and Taylor, 1979). Apparently, this gelatinous matrix may also act as a trap for debris and nutrients (Pedersen, 1990), thus contributing to the overall physical, chemical and biological properties of a mature biofilm. From the viewpoint of microbial physiology, the majority of the biofilm studies in the literature were in the category of accelerated biofilm growth which involved the use of high concentrations of organic nutrients to promote rapid growth. The organic nutrients employed included glucose (La Motta, 1976; Characklis et al., 1982; Gantzer et al., 1988b), glycerol (Gilbert et al., 1989), acetate/citrate (Nielsen, 1987), giucose/peptone (Kuroda et al., 1988) and sucrose/yeast extract (McCoy et al., 1981). These chemostat approaches have provided some useful information on the kinetics of biofilm growth, notably in the design of wastewater treatment facilities. However, the utilization of organic nutrient concentrations that are too high to be realistic for natural biofilms would limit their usefulness for predicting the growth rate of natural biofilm in the real, nutrient-poor, aquatic environment. Perhaps the most valuable contribution of this study to the knowledge of biofilm growth was the elucidation of the origin of the natural biofilm. Many organisms including bacteria, protozoa, fungi and algae have been detected in natural biofilms (McCoy et al., 1981; Pedersen, 1982a, b, 1990). However, their relative importance in the development of a natural biofilm was inconclusive. In this study we provided the first direct biochemical evidence that the origin of natural biofilm was mainly derived from bacterial activity, i.e. bacteria being the major component of biofilms which were established on glass slides with water of natural origin. Based on microscopic observations, Pedersen (1982a) also concluded that algae were not involved in biofilm development in the marine environment. Biofilms contain relatively large quantities of polysaccharides (Characklis et al., 1982) and methods based on the reaction of carbohydrate have been
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3,4-dichlorophenol in a fluvial system. Can. J. Physiol. Pharmac. 62, 971-975. Characklis W. G., Trulear M. G., Bryers J. D. and ZeiverN. (1982) Dynamics of biofilm processes: methods. War. Res. 16, 1207-1216. Costerton J. W., Gessey G. G. and Cheng K. J. (1978) How bacteria stick. Scient. Am. 238, 86-95. Dexter S. C., Sullivan J. D. Jr, Williams J. III and Watson S. W. (1975) Influence of substrate wettability on the attachment of marine bacteria to various surfaces. Appl. Microbioi. 30, 298-308. Gantzer C. J., Kollig H. P., Rittmann B. E. and Lewis D. L. (1988a) Predicting the rate of trace-organic compound removal by natural biofilm. Wat. Res. 22, 191-200. Gantzer C. J., Rittmann B. E. and Herrick E. E. (1988b) Mass transport to streambed biofilm. Wat. Res. 22, 709-722. Gilbert P., Allison D. G., Evans D. J., Handley P. S. and Brown M. R. W. (1989) Growth rate control of adherent bacterial populations. Appl. envir. Microbiol. 55, 1308-1311. Jones G. E., Royle L. G. and Murray L. (1976) The assimilation of trace metal ions by the marine bacteria, Arthrobacter marinus and Pseudomonas cuprodurans. In P,EFERENCT~ 0 Proceedings of the Third International Biodegradation Symposium (Edited by Sharpley J. M. and Kaplan A. M.), Aftrin8 R. P. and Taylor B. F. (1979) Assessment of pp. 889-898. Applied Science, London. microbial fouling in an ocean thermal energy conversion Jordan T. L. and Staley J. T. (1976) Electron microscopy experiment. Appl. envir. Microbiol. 311, 734-739. of succession in the periphyton community of Lake Binder H. (1980) Separation of monosaccharides by highWashington. Microb. Ecol. 2, 241-251. performance liquid chromatography: comparison of Kuroda M., Yuzawa M., Sakakibara Y. and Okamura M. ultraviolet and refractive index detection. J. Chromatogr. (1988) Methanogenic bacteria adhered to solid supports. 189, 414--420. Wat. Pea. 22, 653-656. Blaise C. R. (1986) Micromethod for acute aquatic toxicity assessment using the green alga Selenastrum capricornu- La Motta E. J. (1976) Kinetics of growth and substrate uptake in a biological film system. Appl. etwir. Microbiol. turn. Toxic. Assess. 1, 377-385. 31, 286--293. Borchardt L. G. and Piper C. V. (1970) A gas chromatographic method for carbohydrates as alditol-acetates. Liu D., Lau Y. L., Clmu Y. K. and Pacepavicius G. J. (1992) A simple technique for the determination of bioTappi $3, 257-260. film formation and growth. NWRI Contribution Burkholder J. M. and Wetzel R. G. (1989) Microbial 92-102, National Water Research Institute, Burlington, colonization on natural and artificial macrophytes in Ontario. a phosphorus-limited hardwater lake. J. Phyeol. 25, McCoy W. F., Bryen J. D., Robbins J. and Costerton J. W. 55--65. (1981) Observation of fouling biofilm formation. Can. J. Carey J. H., Fox M. E., Browniec B. G., Metcalfe J. L. and MicrobioL 27, 910-917. Platford R. F. (1984) Disappearance kinetics of 2,4- and
utilized to quantify biofdm mass (Aftring and Taylor, 1979; Characklis et al., 1982). However the physical removal (scrapping or vacuuming) of biofilm from the supporting substratum is required prior to carbohydrate determination for all these procedures. Difficulties have been experienced in the removal of biofilm from the supporting substratum (Aftring and Taylor, 1979). This difficulty is completely eliminated in our direct procedure for the estimation of biofilm mass, thus greatly enhancing the method's simplicity and sensitivity. Moreover, the direct method is very robust and the test can be readily applied to biofilm samples from the marine environment and alkaline lakes. Biofilms can develop on almost any surface exposed to an aqueous environment. Biofilm growth in natural environments is generally slow and our sensitive method can be useful in the estimation of biofilm mass.
Characterization of biofilm development McFeters (3. A. (1984) Biofilm development and its consequences. In Microbial Adhesion and Aggregation (Edited by Marshall K. C.), pp. 109-124, Springer, Berlin. Nielsen P. H. (1987) Biofilm dynamics and kinetics during high-rate sulfate reduction under anaerobic conditions. Appl. envir. Microbiol. 53, 27-32. Pedersen K. (1982a) Method for studying microbial biofilms in flowing-water systems. Appl. envir. Microbiol. 43, 6--13.
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Pedersen K. (1982b) Factors regulating microbial biofilm development in a system with slowly flowing seawater. Appl. envir. Microbioi. 44, 1196-1204. Pedersen K. (1990) Biofilm development on stainless steel and PVC surfaces in drinking water. War. Res. 24, 239-243. Pringle J. H. and Fletcher M. (1983) Influence of substratum wettability on attachment of freshwater bacteria to solid surfaces. Appl. envir. Microbiol. 45, 811-817.