1 CSIRO Land and Water, Floreat, Western Australia
2 Microbiology, School of Biomedical and Chemical Sciences, QEII Medical Centre, The University of Western Australia, Nedlands, Australia
Correspondence
Simon Toze
Simon.Toze{at}csiro.au
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ABSTRACT |
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Abbreviations: ASR, aquifer storage and recovery; CSLM, confocal scanning laser microscopy; D, dilution rate; DAPI, 4',6-diamidine-2'-phenylindole dihydrochloride; GFP, green fluorescent protein; SA, surface area
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INTRODUCTION |
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The attenuation of bacterial pathogens introduced into subsurface environments has been observed in both field and laboratory studies and has been modelled using first-order decay kinetics (Pavelic et al., 1998; Dowd & Pillai, 1997
; Yates & Yates, 1988
). Factors that commonly limit the survival of bacterial pathogens introduced into groundwater include the low level of available nutrients, the lack of oxygen (for obligate aerobes) as well as the competitive, antagonistic and predatory activities of the indigenous microbial population (Gerba & Goyal, 1985
). The practice of aquifer storage and recovery (ASR), which involves the injection of surface water into an aquifer, can create an environment which allows the rapid growth of indigenous and/or introduced bacteria in the aquifer, particularly when the injected water is high in nutrients (Pavelic & Dillon, 1997
; Vecchioli, 1970
). ASR provides a means of storing reclaimed wastewaters prior to re-use for purposes such as irrigation. During an ASR pilot project in South Australia using treated sewage effluent, bacterial growth during injection resulted in biofilm formation within the aquifer matrix immediately surrounding the injection well (Rinck-Pfeiffer, 2000
).
This study was undertaken to determine whether the biofilms developed in an aquifer during ASR could potentially provide a reservoir for pathogenic bacteria. If the recovered water contains a greater number of pathogens than expected, its re-use may pose a potential public health risk. Furthermore, water from the initial stages of recovery is often enriched in dislodged biofilm material and may require special disposal if it contains large numbers of pathogens. In this study, E. coli was used as a representative of the enteric bacterial pathogens and its survival compared to the ubiquitous water-borne micro-organism and opportunistic pathogen Pseudomonas aeruginosa.
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METHODS |
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Quality of water used.
Anaerobic groundwater was collected from the superficial aquifer on the Swan Coastal Plain (Perth, Western Australia) using a submersible electric pump. When required, a sterile 10 % (w/v) peptone (Oxoid) solution in distilled H2O was added to 5 l groundwater to give a final concentration of 0·01 % (v/v). Effluent was collected from a clarification pond, used for sedimentation processing after primary and activated sludge treatment, at the Subiaco Wastewater Treatment Plant, Western Australia. Particulates were removed from the effluent by passing through glass microfibre filter paper (1·2 µm pore size; Whatman). Some chemical properties of the waters used in the reactor experiments are given in Table 1.
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Three coverslips and a water sample were removed from the reactor flasks on days 0, 3, 7, 10, 15 and 21 during the E. coli experiment and on days 0, 7, 14, 26 and 40 during the P. aeruginosa experiment. Coverslip samples were immediately rinsed three times in PBS (24 g NaCl l-1; 0·6 g KCl l-1; 4·32 g Na2HPO4 l-1; 0·72 g KH2PO4 l-1 at pH 7·0) to remove unattached or loosely adhered cells prior to further processing (described below). Viable E. coli numbers in the outflow from each flask were monitored during the E. coli experiment. At the end of the P. aeruginosa experiment (day 47), each flask that had been inoculated with P. aeruginosa was destructively sampled to determine the extent of biofilm development and numbers of attached P. aeruginosa on the remaining coverslips, the coverslip holder and wall of each reactor flask.
Culturable cell counts.
Coverslip samples were placed in 12 ml PBS immediately after rinsing. Cells were removed from the surface by scraping with a sterile disposable cell scraper (17 mm blade length; Sarstedt), followed by sonication in a 50 kHz water-bath (10 min) and vortexing (1 min). The same procedure was followed to remove cells, after rinsing, from the coverslip holders and reactor flasks at the end of the P. aeruginosa experiment, using 20 ml or 100 ml volumes of PBS, respectively. Culturable counts of GFP-labelled cells in water or processed biofilm samples were performed by either the drop-on-plate method (with six replicate 10 µl drops of the appropriate dilutions) or spread-plate method (triplicate plates using undiluted 100 µl or 200 µl volumes) using LB agar containing the appropriate antibiotic. The number of colonies showing green fluorescence under blue light illumination was recorded.
Epifluorescence microscopy.
For determination of fluorescent cell numbers, aqueous samples were filtered onto black polycarbonate filters (0·2 µm pore size, 25 mm diameter; Millipore) and mounted on glass slides as described by Hobbie et al. (1977). Coverslip samples were air-dried after rinsing and mounted on a glass slide using nail varnish. Mounted coverslips were overlaid with mineral oil and a clean coverslip before viewing. Filters and coverslip samples were viewed under oil immersion with a 100x times; Plan objective on a Leitz Diaplan microscope fitted with a Leitz Pleomopak fluorescence attachment. GFP-conferred fluorescence was visualized under illumination with blue light (excitation 450490 nm; suppression 515 nm). The number of cells in a minimum of 20 randomly chosen fields of view was determined for each filtered aqueous sample and each coverslip sample.
The SA coverage of the biofilms was determined by staining the coverslip samples with the nucleic acid stain DAPI (4',6-diamidine-2'-phenylindole dihydrochloride) at a concentration of 0·5 µg ml-1 for 10 min in the dark. Ten photomicrographs from each coverslip were taken under UV light (excitation 340380 nm; suppression 430 nm) at 1000x times; magnification with a cooled slow scan PXL CCD camera (Photometrics). The area of DAPI-conferred fluorescence (in µm2) was measured in each photomicrograph using IPLab Spectrum version 3 (Scanalytics). The SA values were converted to a percentage of the photomicrograph area, which was 6143 µm2, and the mean value for each coverslip sample determined.
Confocal scanning laser microscopy (CSLM).
After rinsing, coverslip samples were mounted in screw-top circular stainless steel chambers with a viewing area of 7·5 mm radius. The chambers were filled with PBS to prevent the biofilm drying during viewing and the underside of each coverslip was cleaned with 70 % ethanol. Biofilms were viewed using a Bio-Rad MRC1000/1024 UV confocal scanning laser microscope, mounted on a Nikon Diaphot 300 with a Nikon 60x times; water-immersion PlanApo objective lens (numerical aperture of 1·2). A correction collar allowed adjustment for the thickness of the coverslip. The microscope was controlled by the COMOS software (Bio-Rad Microscience) and images were collected using a Kalman mathematical filter (where n=3) to reduce background noise. All images were collected using a confocal pinhole size of 2·5 mm. For GFP detection, 488 nm argon laser light was used to excite the specimen and fluorescence was collected through a 522/35 nm emission filter. Power and gain settings were chosen to supply the most sensitivity without picking up autofluorescence in the negative control biofilms (i.e. from flasks that were not inoculated with GFP-labelled cells). A z-axis stepping motor, which allows movement of the focal plane in precise increments (minimum of 0·1 µm), was used to collect a z-series of images for each field of view. Confocal microscope images were processed using Confocal Assistant Software version 4.02 (Bio-Rad).
Data analysis.
Removal rates were determined by calculating the slope and correlation coefficient (r2) of the linear regression of log-transformed cell concentration data according to the first-order decay relation given by: C=C010-kt where C is the microbial concentration at time t, C0 the initial concentration on day 0 and k the removal rate. The concentration data were mean values from replicate culture-based or direct cell counts from a single water sample or coverslip sample. Where reactor flasks were run in duplicate (i.e. test flasks containing coverslips), the removal rate associated with each flask was calculated first and then the mean value and standard deviation of the two rates determined. The cell removal rates derived from direct GFP-fluorescent cell counts are referred to as total and those from GFP-fluorescent colony counts are referred to as viable. A Student's t-test (one-tailed distribution for two samples with unequal variance) was used to determine if differences between test flasks with flow-through of the different water types were significant.
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RESULTS |
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The GFP plasmids were found to be stable in the type strains of E. coli and P. aeruginosa over 40 generations of growth in LB broth without the addition of antibiotic when incubated at 28 °C in a static incubator (data not shown). During incubation of E. coli(pEGFP) in sterile groundwater microcosms for 50 days, the number of c.f.u. on LB with or without antibiotic did not significantly differ (P>0·05), demonstrating maintenance of plasmid expression in the culturable cell population. In addition, no significant differences between the total number of cells (enumerated using the nucleic acid stain DAPI) and the number of GFP-fluorescent cells were detected during this time (Fig. 2a). Incubation of E. coli(pEGFP) in sterile effluent microcosms gave similar results (data not shown). Maintenance of fluorescence in P. aeruginosa(pSMC21) in sterile groundwater microcosms was also demonstrated in the culturable cell population although the proportion of the total cell population with GFP-conferred fluorescence decreased over time (Fig. 2b
). These results are in agreement with previous studies demonstrating that intact cells may maintain GFP-conferred fluorescence even if non-viable but that some intact dead or dying cells may lose GFP-conferred fluorescence under conditions of stress such as nutrient limitation (Banning et al., 2002
) or heat (Lowder et al., 2000
). It is likely that this loss of fluorescence is linked to leakage of the protein though a damaged cell membrane. Although the GFP plasmidhost constructs used in this study displayed different patterns of fluorescence loss, both GFP plasmids were considered as suitable cellular markers (but not viability indicators) for use under non-selective, anaerobic, nutrient-limited conditions.
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Removal rates of GFP-labelled cells
The rates of removal of total and viable E. coli(pEGFP) and P. aeruginosa(pSMC21) cells from biofilms and from the water phase are shown in Table 2. The time periods of days 010 for E. coli and 014 for P. aeruginosa were chosen for determining removal rates as cell numbers in all samples between these periods were above detection limits. Persistence of GFP-labelled cells inoculated into the reactor was a result of the interplay between the dilution or washout effect of continuous flow, cell growth and death rates and attachment/detachment processes. It was not possible to confirm whether any of the viable inoculated cells or their progeny had lost the GFP plasmid or expression of the plasmid during the reactor experiments due to the presence of the mixed microbial population. However, the results from the sterile microcosms, discussed above, suggest that this is unlikely to have occurred over the 21 or 40 day time period.
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For P. aeruginosa, there was no significant difference between the removal rates of attached cells in flasks with groundwater or effluent flow-through and both rates were slower than that measured for E. coli. There was also little difference in removal rates (total or viable) of planktonic P. aeruginosa between control and test flasks. However, there was a pronounced difference between the behaviour of planktonic P. aeruginosa in flasks with flow-through of groundwater and those with effluent. After day 14 the number of viable P. aeruginosa in the flasks with groundwater flow-through fell below detection limits whereas there were still detectable levels in the flasks with effluent flow-through by day 40. The rate of planktonic P. aeruginosa removal in flasks with effluent flow-through measured between days 0 and 40 was -0·12 log10 c.f.u. ml-1 day-1, with or without biofilms, which was slower than the theoretical washout rate.
Prolonged persistence of P. aeruginosa compared to E. coli
A comparison of the length of time taken for the GFP-labelled cells in reactor flasks to fall below the detection limits also reveals that the most pronounced difference between E. coli and P. aeruginosa behaviour occurred in flasks with effluent flow-through (Fig. 5). In these flasks, P. aeruginosa persisted in an attached state and in the water phase for longer than E. coli and for longer than the theoretical time for cells to fall below the detection limits as a result of dilution.
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DISCUSSION |
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Established biofilms developed from indigenous river water bacteria have been shown previously to reduce persistence of introduced E. coli and other enteric pathogens (Camper et al., 1985). Furthermore, changing biofilm dynamics and pathogen persistence as a result of an increase in nutrient levels has been reported elsewhere. The survival of a C. jejuni strain in heterogeneous tap-water biofilms was shown to be significantly reduced by the addition of serine, a carbon source known to be favoured by C. jejuni, during which time the number of indigenous biofilm microflora increased (Buswell et al., 1998
, 1999
). These studies demonstrate that under certain conditions biofilms may represent sites of intensified competition for limiting nutrients.
By comparison, P. aeruginosa persisted in the biofilms in the flasks with effluent flow-through for much longer than E. coli. In both the test flasks and the control flask with effluent flow-through, the rate of P. aeruginosa removal from the water phase slowed dramatically after day 14 to a rate slower than the theoretical washout rate, suggesting that P. aeruginosa was growing in the treated effluent. It is unlikely that there was enough oxygen available in the airtight reactor flasks to sustain growth of P. aeruginosa, which is an obligate aerobe. Although the mean concentration of dissolved oxygen in the treated effluent was higher than the groundwater upon collection (Table 1), the oxygen levels in the water decreased during the course of the experiment due to constant bubbling of water in the holding reservoirs with nitrogen gas. Thus, growth of P. aeruginosa most likely occurred through utilization of nitrate as the terminal electron acceptor, which was present at much higher concentrations in the effluent than the groundwater (Table 1
). P. aeruginosa also has the ability to utilize a wider range of organic molecules as carbon and energy sources compared to Enterobacteriaceae (Bergey et al., 1984
). As there was no difference in P. aeruginosa removal rates between the control and test flasks, the increased persistence of P. aeruginosa in effluent could not be attributed to an interaction with the biofilms.
The laboratory reactor used in this study was not a perfect simulation of aquifer conditions during ASR. Thus, survival times of the micro-organisms reported here may vary considerably from survival times in the field. Nonetheless, this study has shown that addition of a high-nutrient water to a low-nutrient environment may stimulate biofilm development but have a detrimental effect on survival of bacteria such as E. coli. Thus, the biofilm development which has been found to occur during ASR using reclaimed sewage effluent (Rinck-Pfeiffer, 2000) may not pose a health risk with respect to the persistence of enteric bacterial pathogens in the aquifer during storage of the water. On the other hand, growth of the opportunistic pathogen P. aeruginosa, which is able to compete more effectively with the indigenous microbial population for available nutrients, may occur during ASR.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Bergey, D. H., Holt, J. G. & Kreig, N. R. (1984). Bergey's Manual of Systematic Bacteriology, vol. 1. Baltimore: Williams & Wilkins.
Block, J. C. (1992). Biofilms in drinking water distribution systems. In Biofilms Science and Technology, pp. 469485. Edited by T. R. Bott, L. Melo, M. Fletcher & B. Capedeville. Dordrecht: Kluwer.
Bloemberg, G. V., O'Toole, G. A., Lutenberg, B. J. J. & Kolter, R. (1997). Green fluorescent protein as a marker for Pseudomonas spp. Appl Environ Microbiol 63, 45434551.[Abstract]
Buswell, C. M., Herlihy, Y. M., Lawrence, L. M., McGuiggan, J. T. M., Marsh, P. D., Keevil, W. & Leach, S. A. (1998). Extended survival and persistence of Campylobacter spp. in water and aquatic biofilms and their detection by immunofluorescent-antibody and -rRNA staining. Appl Environ Microbiol 64, 733741.
Buswell, C. M., Herlihy, Y. M., Keevil, C. W., March, P. D. & Leach, S. A. (1999). Carbon load in aquatic ecosystems affects the diversity and biomass of water biofilm consortia and the persistence of the pathogen Campylobacter jejuni within them. J Appl Microbiol 85, 161S167S.
Buswell, C. M., Nicholl, H. S. & Walker, J. T. (2001). Use of continuous culture bioreactors for the study of pathogens such as Campylobacter jejuni and Escherichia coli O157 in biofilms. Methods Enzymol 337, 7078.[Medline]
Camper, A. K, LeChevallier. M. W., Broadaway, S. C. & McFeters, G. A. (1985). Growth and persistence of pathogens on granular activated carbon filters. Appl Environ Microbiol 50, 13781382.[Medline]
Camper, A. K., Jones, W. L. & Hayes, J. T. (1996). Effect of growth conditions and substratum composition on the persistence of coliforms in mixed-population biofilms. Appl Environ Microbiol 62, 40144018.[Abstract]
Colbourne, J. S., Pratt, D. J., Smith, M. G., Fisher-Hoch, S. P. & Harper, D. (1984). Water fittings as sources of Legionella pneumophila in a hospital plumbing system. Lancet i, 210213.
Cormack, B. P., Valdivia, R. H. & Falkow, S. (1996). FACS-optimized mutants of the green fluorescent protein (GFP). Gene 173, 3338.[CrossRef][Medline]
Costerton, J. W., Lewandowski, Z., Caldwell, D. E., Korber, D. R. & Lappin-Scott, H. M. (1995). Microbial biofilms. Annu Rev Microbiol 49, 711745.[CrossRef][Medline]
Decho, A. W. (2000). Microbial biofilms in intertidal systems: an overview. Continental Shelf Res 20, 12571273.[CrossRef]
Dowd, S. E. & Pillai, S. D. (1997). Survival and transport of selected bacterial pathogens and indicator viruses under sandy aquifer conditions. J Environ Sci Health 32, 22452258.
Gerba, C. P. & Goyal, S. M. (1985). Pathogen removal from wastewater during groundwater recharge. In Artificial Recharge of Groundwater, pp. 283317. Edited by T. Asano. Boston, MA: Butterworth.
Gilbert, P. & Brown, M. R. W. (1995). Mechanisms of the protection of bacterial biofilms from antimicrobial agents. In Microbial Biofilms, pp. 118130. Edited by H. M. Lappin-Scott & J. W. Costerton. Cambridge: Cambridge University Press.
Hobbie, J. E., Daley, R. J. & Jasper, S. (1977). Use of nucleopore filters for counting bacteria by fluorescence microscopy. Appl Environ Microbiol 33, 12251228.[Medline]
LeChevallier, M. W., Babcock, T. M. & Lee, R. G. (1987). Examination and characterization of distribution system biofilms. Appl Environ Microbiol 53, 27142724.[Medline]
Lowder, M., Unge, A., Maraha, N., Jansson, J. K., Swiggett, J. & Oliver, J. D. (2000). Effect of starvation and the viable-but-nonculturable state on green fluorescent protein (GFP) fluorescence in GFP-tagged Pseudomonas fluorescens A506. Appl Environ Microbiol 66, 31603165.
Mackay, W. G., Gribbon, L. T., Barer, M. R. & Reid, D. C. (1999). Biofilms in drinking water systems: a possible reservoir for Helicobacter pylori. J Appl Microbiol 85, 52S59S.
Marrao, G., Verissimo, A., Bowker, R. G. & daCosta, M. S. (1993). Biofilms as major sources of Legionella spp. in hydrothermal areas and their dispersion into stream water. FEMS Microbiol Ecol 12, 2533.[CrossRef]
Momba, M. N. B., Cloete, T. E., Venter, S. N. & Kfir, R. (1999). Examination of the behaviour of Escherichia coli in biofilms established in laboratory-scale units receiving chlorinated and chloraminated water. Water Res 33, 29372940.[CrossRef]
Murga, R., Forster, T. S., Brown, E., Pruckler, J. M., Fields, B. S. & Donlan, R. M. (2001). Role of biofilms in the survival of Legionella pneumophila in a model potable-water system. Microbiology 147, 31213126.
Murgel, G. A., Lion, L. W., Acheson, C., Shuler, M. L., Emerson, D. & Ghiorse, W. C. (1991). Experimental apparatus for selection of adherent microorganisms under stringent growth conditions. Appl Environ Microbiol 57, 19871996.[Medline]
Olofsson, A.-C., Zita, A. & Hermansson, M. (1998). Floc stability and adhesion of green-fluorescent-protein-marked bacteria to flocs in activated sludge. Microbiology 144, 519528.[Abstract]
Pavelic, P. & Dillon, P. J. (1997). Review of international experience in injecting natural and reclaimed waters into aquifers for storage and reuse. Centre for Groundwater Studies Report No. 74, South Australia.
Pavelic, P., Dillon, P. J., Barry, K. E. & Herczeg, A. L. (1998). Well clogging effects determined from mass balances and hydraulic response at a stormwater ASR site. In Artificial Recharge of Groundwater: Proceedings of the Third International Symposium on Artificial Recharge of Groundwater, pp. 6167. Edited by J. H. Peters. Amsterdam: Balkema.
Rinck-Pfeiffer, S. (2000). Physical and biochemical clogging processes arising from aquifer storage and recovery (ASR) with treated wastewater. PhD thesis, Flinders University of South Australia.
Robinson, P. J., Walker, J. T., Keevil, C. W. & Cole, J. (1995). Reporter genes and fluorescent probes for studying the colonisation of biofilms in a drinking water supply line by enteric bacteria. FEMS Microbiol Lett 129, 183188.[CrossRef][Medline]
Rogers, J., Dowsett, A. B., Dennis, P. J., Lee, J. V. & Keevil, C. W. (1994). Influence of plumbing materials on biofilm formation and growth of Legionella pneumophila in potable water systems. Appl Environ Microbiol 60, 18421851.[Abstract]
Scott, K. P., Mercer, D. K., Glover, L. A. & Flint, H. J. (1998). The green fluorescent protein as a visible marker for lactic acid bacteria in complex ecosystems. FEMS Microbiol Ecol 26, 219230.[CrossRef]
Szewzyk, U., Szewzyk, R., Manz, W. & Schleifer, K.-H. (2000). Microbiological safety of drinking water. Annu Rev Microbiol 54, 81127.[CrossRef][Medline]
Van der Wende, E., Characklis, W. G. & Smith, D. B. (1989). Biofilms and bacterial drinking water quality. Water Res 23, 13131322.[CrossRef]
vanLoosdrecht, M. C. M., Lyklema, J. L., Norde, W. & Zehnder, A. J. B. (1990). Influence of interfaces on microbial activity. Microbiol Rev 54, 7587.
Vecchioli, J. (1970). A note on bacterial growth around a recharge well at Bay Park, Long Island, New York. Water Resour Res 6, 14151419.
Vess, R. W., Anderson, R. L., Carr, J. H., Bond, W. W. & Favero, M. S. (1993). The colonization of solid PVC surfaces and the acquisition of resistance to germicides by water micro-organisms. J Appl Bacteriol 74, 215221.[Medline]
Walker, J. T., Mackerness, C. W., Rogers, J. & Keevil, C. W. (1995). Heterogeneous mosaic biofilm a haven for waterborne pathogens. In Microbial Biofilms, pp. 196204. Edited by H. M. Lappin-Scott & J. W. Costerton. Cambridge: Cambridge University Press.
Yates, M. V. & Yates, S. R. (1988). Virus survival and transport in ground water. Water Sci Technol 20, 301307.
Received 8 August 2002;
revised 7 October 2002;
accepted 8 October 2002.