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Applied and Environmental Microbiology, December 2008, p. 7204-7210, Vol. 74, No. 23
0099-2240/08/$08.00+0     doi:10.1128/AEM.00341-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Evidence for Growth of Enterococci in Municipal Oxidation Ponds, Obtained Using Antibiotic Resistance Analysis

Elaine Moriarty, Fariba Nourozi, Beth Robson, David Wood, and Brent Gilpin^*

Institute of Environmental Science & Research Limited, Christchurch Science Centre, 27 Creyke Road, P.O. Box 29-181, Christchurch, New Zealand

Received 10 February 2008/ Accepted 14 September 2008

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The Christchurch wastewater treatment plant uses a series of six oxidation ponds to reduce the bacterial load of treated effluent before it is discharged into the local estuary. To ensure that this discharge does not adversely affect water quality in the receiving environment, local regulations specify maximum levels in the discharge for a number of parameters, including enterococci. Between 2001 and 2006, regulations required fewer than 300 enterococci per 100 ml in summer. During this period, the discharge intermittently exceeded this limit, with unexplained levels of enterococci of up to 180,000/100 ml. Characterization of these enterococci by antibiotic resistance analysis showed that enterococci sampled over 4 months had almost identical resistance profiles. In contrast, enterococci from raw sewage and wildfowl from around the oxidation ponds had a diverse range of antibiotic resistance profiles that could be distinguished from each other and also from those of enterococci from the discharge. The hypothesis of a clonal nature of the enterococci in the discharge was supported by molecular genotype analysis, suggesting that these bacteria may have replicated in the pond environment rather than being reflective of breakthrough in the sewage treatment process or the result of recent wildfowl inputs to the ponds. This study highlights the usefulness of antibiotic resistance analysis in identifying this phenomenon and is the first report of apparent replication of a specific type of enterococci in an oxidation pond environment.

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Each day, 150 million liters of wastewater is treated at the Christchurch wastewater treatment plant (WWTP) on the South Island of New Zealand. Primary sedimentation tanks, trickling filter towers, solids contact, aeration tanks, and clarifying tanks remove most of the oxygen demand and suspended solids and some of the bacteria before, over a period of 3 weeks, the effluent passes through a series of six oxidation ponds, encompassing 230 ha (Fig. 1). Within the oxidation ponds, the sun's UV radiation and grazing by zooplankton provide an inhospitable environment for bacteria present in the effluent, resulting in the removal of an estimated 99.999% of fecal coliforms. Twice daily, for the first few hours following high tide, treated output from pond 6 is discharged into the estuary of the Avon and Heathcote rivers. Water in pond 6 is monitored twice weekly for a range of water quality indicators, including fecal coliforms and enterococci. These indicator bacteria are ubiquitous in the raw sewage entering the plant. While they are usually not harmful themselves, they indicate the possible presence of pathogenic bacteria (Campylobacter, Salmonella, etc.), viruses (norovirus, hepatitis A virus, etc.), and protozoans (Cryptosporidium, Giardia, etc.) that can be found in the intestinal tracts of human and animals. Maximum acceptable levels of each of the measured parameters have been established by the Regional Council to ensure safe bathing in the receiving waters of the discharge estuary. For the microbial indicators, separate summer (November to February) and winter (March to October) limits have been established. The discharge consent required that no more than six of eight consecutive samples taken were to exceed the allowable limit for enterococci of a most probable number of 300/100 ml in summer and 1,500/100 ml in winter. In addition, no more than two of eight consecutive samples were to exceed the higher value of 500/100 ml in summer and 2,000/100 ml in winter. The discharge conditions also specified values for fecal coliforms, for which the lower values were 2,000/100 ml in summer and 10,000/100 ml in winter, with higher values of 20,000/100 ml in both summer and winter. The limits are higher in winter to reflect the reduced effectiveness of the oxidation ponds under lower UV conditions.

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FIG. 1. Layout of oxidation ponds at Christchurch WWTP (http://www.ccc.govt.nz/WasteWater/TreatmentPlant). Arrows indicate the direction of flow through ponds 1 to 6. Image courtesy of Terralink. Copyright 2001, Terralink International Limited.

Between December 2005 and May 2006, unusually high levels of enterococci were detected in pond 6 immediately prior to discharge into the estuary (Fig. 2). Interestingly, these peaks occurred at a time when fecal coliform levels were within the normal range. The question was posed as to whether these "exceedence" enterococci were breakthrough from the treatment process (thereby indicative of human sewage and a significant health risk), whether they were from the wildfowl that occupied the ponds, or whether they were from some other source. The oxidation ponds are part of the 390-ha Te Huingi Manu wildlife reserve, with up to 30,000 ducks, geese, and other wildfowl inhabiting the ponds. Access to the ponds is restricted by a series of gates and fences which prevent access by any other large animals.

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FIG. 2. Levels of fecal coliforms and enterococci in oxidation pond 6 from November 2004 to February 2007. Upper enterococcal consent limits are indicated by solid lines. Fecal coliform upper consent limits of 20,000/100 ml were not observed. Filled circles indicate the samples on which ARA was performed. MPN, most probable number.

A range of tools have been developed for clarifying the potential sources of fecal indicator bacteria, including the use of fluorescent whitening agents, fecal sterols, molecular markers, and antibiotic resistance analysis (ARA), among others (8, 9, 11, 24, 28). The ARA method was first published in 1996 and has been used to estimate the sources of pollution in various environmental settings (13, 26, 28, 29). This method has been applied to both Escherichia coli and enterococci and involves collecting isolates from potential fecal or effluent sources. The profiles of the resistance of these bacteria to several concentrations of up to 12 antibiotics form a library of antibiotic resistance profiles of known isolates that unknown samples can be compared with.

The initial hypothesis of this study was that the high levels of enterococci in the oxidation pond discharge were due to wildfowl inputs from the considerable population of birds that inhabit the ponds. A second hypothesis was that the enterococci were breakthrough from the sewage treatment process. In this study, an ARA library of isolates from potential sources including wildfowl and raw sewage samples was established and used for comparison with the enterococcal isolates from pond 6 in an attempt to determine the source of these enterococci.

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Sample collection and isolation of enterococci.
Samples (100 ml) from final-stage oxidation pond 6 of the Christchurch WWTP were analyzed twice weekly between November 2004 and February 2007 for enterococci by using Enterolert assays (Idexx Laboratories Inc., Westbrook, ME) and for fecal coliforms by the Colilert system (Idexx Laboratories Inc.). Samples were stored at 4°C, and when high levels of enterococci were detected by the WWTP laboratory, they were confirmed by filtering serially diluted samples through 0.45-µm filters (type GN-6; Gelman Sciences) within 36 h of the original sample collection. The filters were transferred to a 50-mm petri dish containing M-Ent agar (Merck Inc.) and incubated at 37°C for 48 h. For comparison, additional samples were collected from primary screened raw wastewater as it entered the plant (n = 14 samples). Fresh fecal samples were collected from the shores of the oxidation ponds at the treatment plant and also from a town park approximately 10 km away. Wildfowl samples included scats from Canadian geese (Branta canadensis, n = 12), ducks (Anas spp., n = 10), swans (Cygnus spp., n = 9), and 20 other unidentified wildfowl birds. For each of these samples, a 10% (wt/vol) fecal slurry in saline buffer (0.15 M NaCl, 0.002 M KH₂PO₄, pH 7.3) was prepared and processed as described above.

Attributes of pond 6 were measured allowing the calculation of conductivity, biological oxygen demand, dissolved oxygen, percent O₂ saturation, ammonia, nitrite, nitrate, pH, reactive and total phosphorus, suspended solids, percent transmission, water and air temperatures, rainfall, and wind speed.

Characterization of enterococci.
Isolates were characterized as described by Wiggins (28). Clearly separated, individual colonies from M-Ent plates were transferred by using sterile wooden toothpicks into microwell plates (Sarstedt, Germany) containing 0.2 ml of Enterococcosel broth (Oxoid, United Kingdom). Plates were sealed and incubated for 48 h at 35°C. Isolates that demonstrated the ability to hydrolyze esculin through the formation of black pigment in the wells were further characterized. Approximately 3% of the isolates chosen at random were further characterized by testing for the production of catalase, Gram stain reaction, and growth in various carbohydrates (1). Selected Enterococcus isolates were identified to the species level by partial sequencing of the RNA polymerase alpha subunit (rpoA) and phenylalanyl-tRNA synthase (pheS) genes (20). Bacterial cells were suspended in distilled water and heated at 95°C for 5 min to release their DNA. Amplitaq polymerase (Applied Biosystems) was used to amplify 474-bp pheS and 810-bp rpoA PCR products as described by Naser et al. (20). PCR products were sequenced on an ABI 3700 sequencer, and sequenced DNA was compared with sequences in GenBank by using the BLAST algorithm (18).

ARA.
The antibiotic resistance profiles of the isolates were determined as previously described (28). Enterococcal isolates were transferred to a set of tryptic soy agar plates (Merck) containing different antibiotics (Sigma-Aldrich), each at three or four concentrations (Table 1), and a control plate containing no antibiotics. Plates were incubated for 48 h at 37°C, and the highest concentration of antibiotic which permitted visible growth was recorded. Isolates that did not grow on the control plates were not considered in the analyses. Each ARA type consisted of a single resistance pattern with no allowance for variability within a designated type.

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TABLE 1. Antibiotics used for resistance analysis

PFGE.
A selection of isolates (n = 27) were analyzed by pulsed-field gel electrophoresis (PFGE). Isolates were grown on Columbia blood agar plates, and agarose plugs were prepared by the PulseNet Listeria protocol (10). Plugs containing lysed enterococcal cells were digested with 40 U of SmaI restriction enzyme for 3 to 4 h at 25°C. Electrophoresis was performed with switch times of 5 to 35 s at 6 V/cm and at 14°C for 18 h through 1% SeakemGold agarose gel with a CHEF DRIII electrophoresis unit (Bio-Rad Laboratories). Each gel included the PulseNet universal size standard Salmonella Braenderup H9812 digested with XbaI (14) and a laboratory strain of Enterococcus faecalis digested with SmaI as a quality control to ensure that complete digestion occurred. Isolates were analyzed in BioNumerics 5.1 (Applied Maths, Belgium) by using the Dice coefficient, the unweighted-pair group method using average linkages, an optimization of 1.0%, and a tolerance of 1.5%.

Statistical analysis.
The diversity index (DI) (23) for various sources was estimated, at the 95% confidence interval, by the approach of Grundmann et al. (12). Discriminant analysis (DA) was undertaken with XLSTAT 2007.4 (www.xlstat.com) and profiles of resistance to all of the antibiotics tested, except bacitracin (for which some data items were missing) and vancomycin, which lacked any useful discriminatory power. Samples were assigned at random to either a library or a validation group, and by using DA, the validation set was tested against the library data.

For comparison of the physical and chemical attributes of the pond, samples were divided into a low enterococcal count group (<300/100 ml, n = 127) and a high enterococcal count group (>1,000/100 ml, n = 48). Student's t test was used to compare the means between these groups, and a significance value of 0.01 was used.

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Recovery of enterococci from oxidation pond, sewage, and fecal samples.
Fecal coliform levels in the discharge from pond 6 ranged between 45 and 1,720/100 ml, with a median of 342/100 ml. At no point did fecal coliform levels exceed the summer or winter higher value discharge conditions (Fig. 2). In contrast, enterococcal levels ranged from 5 to 182,200 CFU/100 ml, with 20% of the samples exceeding the higher discharge conditions of 500 CFU/100 ml for summer samples and 2,000 CFU/100 ml for winter samples. From 12 of these exceedences (indicated by a filled circle in Fig. 2), 440 enterococci were isolated in groups of between 10 and 80 isolates from each sampling occasion. From the 15 raw sewage samples collected from 30 May 2006 to 27 November 2006, between 10 and 77 enterococcal isolates were recovered per sample, providing a total of 517 sewage isolates for further analysis.

Fifty wildfowl scats were analyzed for the presence of fecal coliforms, E. coli, and enterococci. Of these, 24 scats contained no detectable enterococci or fecal coliforms, including 83% of those from Canadian geese (n = 12), 30% of those from ducks (n = 10), 55% of those from swans (n = 9), and 25% of those from unidentified wildfowl (n = 20). The levels of fecal coliforms, E. coli, and enterococci in the remaining samples are listed in Table 2. Excluding scats with levels of indicators below the detection limit, an average of 1.37 x 10⁶ g^–1 fecal coliforms (median, 5.0 x 10³ g^–1), 8.83 x 10⁵ g^–1 (median, 5.0 x 10³ g^–1) E. coli cells, and 7.72 x 10⁴ g^–1(median, 2.0 x 10³ g^–1) enterococci were found in each scat. Enterococcal antibiotic resistance profiles were generated from 18 of the 26 scats collected.

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TABLE 2. Enumeration of fecal coliforms, E. coli bacteria, and enterococci in wildfowl scats

More than 95% of the enterococcal isolates were esculin positive and catalase negative. Fifty randomly selected isolates were characterized further and in all cases were gram-positive cocci that grew in 6.5% NaCl and fermented mannitol, sucrose, and lactose, consistent with classification as enterococci. More than 80% of these isolates fermented sorbitol and did not ferment arabinose, which identified them as E. faecalis. Sequencing of the PCR-amplified rpoA and pheS genes of 11 isolates from the oxidation ponds identified all of these as E. faecalis, while sequencing of isolates from raw sewage identified 4 as E. faecalis and 3 as Enterococcus faecium.

Resistance patterns of enterococci.
Antibiotic resistance profiles were determined for a total of 1,484 isolates. The isolates from both raw sewage and wildfowl displayed a wide variety of resistance patterns. Four hundred seventy-six different resistance patterns were observed among the 517 raw sewage isolates examined (DI = 99.9% ± 0.03%). The most common profile seen (n = 7) was that of no resistance to any antibiotic, while the second most common profile (n = 5) demonstrated only resistance to kanamycin (KAN; 30 mg/ml). Only one isolate was resistant to vancomycin (30 mg/ml).

Between 8 and 80 colonies were analyzed from each of 18 wildfowl scats, resulting in 527 isolates with 271 distinct antibiotic resistance profiles (DI = 99.2% ± 0.16%). Of these, 184 profiles were observed only once, while the two most frequently observed patterns were observed 18 times each. Within each scat, the DIs ranged from 91.1 to 100%, with up to 34 different profiles recovered from an individual scat (mean, 19).

In stark contrast, of the 440 isolates from the high-level oxidation pond discharges, only 32 different antibiotic resistance profiles were identified (DI = 79.6% ± 2.48%). Table 3 illustrates the variations in antibiotic profile observed. Of the 80 isolates analyzed from the sample taken on 16 February 2006, 79 had the same antibiotic resistance profile, with the one different isolate differing only by resistance to 10 µg ml^–1 bacitracin. Samples taken on 17 (n = 80), 21 (n = 40), and 27 (n = 40) February were all indistinguishable by ARA and differed from the previous dominant pattern only by resistance to 20 µg ml^–1 chlortetracycline hydrochloride. Minor variations in antibiotic resistance profile were observed in samples taken in March and April. Samples taken on 1, 5, and 11 May yielded 76 isolates with the same profile, and this was identical to the dominant pattern observed more than 2 months earlier on 17 to 27 February (pattern shown in Table 3).

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TABLE 3. Percentages of enterococcal isolates from oxidation pond 6 with resistance to antibiotics at various concentrations tested

PFGE analysis of isolates.
Eleven isolates from pond 6 with dominant ARA profiles were analyzed by PFGE. Three of these were collected on 13 March, seven were collected on 1 May, and one was collected on 4 May. The March isolates displayed minor variations in resistance to three of the antibiotics tested, but all of these pond 6 isolates were indistinguishable by PFGE analysis (Fig. 3). In contrast, 16 other isolates from raw sewage all had different SmaI PFGE patterns.

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FIG. 3. PFGE patterns of enterococci. Lanes 1, 8, and 15, Salmonella Braenderup H9812 size standard (bands sizes are marked in kilobases); lanes 2 to 7 and 9, selection of isolates from untreated sewage; lanes 10 to 13, oxidation pond 6 enterococcal isolates from 1 May, 4 May, 1 May, and 13 March, respectively; lane 14, E. faecalis isolate.

Statistical analysis.
A comparison of antibiotic resistance profiles of bacteria from wildfowl, sewage, and oxidation ponds was undertaken by three different approaches. Chi-square analysis of the prevalence of resistance to antibiotics among enterococci from each source (Table 4) indicated that the overall resistance pattern of isolates from oxidation pond 6 was not the same as that of bacteria from either of the sources sampled (P < 0.0001). The DI of the oxidation pond isolates (79.59% ± 2.48%) was also significantly different (P < 0.0001) from that of the sewage (99.93% ± 0.01%) and wildfowl groups (99.24% ± 0.16%). For the third analysis, the samples were assigned at random to one of two groups, resulting in a library data set containing 822 isolates and a validation set containing 662 isolates. By DA, the rate of correct classification ranged from 55% for humans to 94% for the oxidation ponds, which gave an average rate of correct classification for the individual isolates of 73% (Table 5). This correct rate of classification is significantly greater than would be achieved by random assignment of the isolates (33%). Although some of the isolates were incorrectly assigned, this analysis demonstrates that correct rates of isolation well exceeded those of misclassification. Similar results and rates of correct classification have been reported by other researchers (28, 29).

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TABLE 4. Percentage of isolates from each source with resistance to the specified antibiotic

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TABLE 5. DAa of antibiotic resistance profiles of enterococci for the validation set

Comparison of the means of physical, chemical, and meteorological attributes at pond 6 by using Student's t test found that samples with enterococcal levels above 1,000/100 ml had higher levels of ammonia (32.7 versus 30.6 ppm), reactive phosphorus (5.9 versus 5.4 ppm), total phosphorus (7.0 versus 6.4 ppm), suspended solids (25.7 versus 17 ppm), and total Kjeldahl N (37 versus 34) and a higher water temperature (17.0 versus 13.7°C). These samples had a lower measured 5-day biological oxygen demand (the amount of dissolved oxygen consumed in 5 days by biological processes breaking down organic matter) (14.2 versus 17.3 ppm), a lower nitrate level (0.1 versus 0.15 ppm), and a lower percent transmission (40 versus 44%). The other measured parameters, including recent rainfall and wind, were not significantly different.

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One hypothesis of this study was that the high levels of enterococci in the oxidation pond discharge tested were due to wildfowl inputs from the considerable population of birds that inhabit the ponds. A second hypothesis was that the enterococci were breakthrough from the sewage treatment process. As a first strategy to address the wildfowl possibility, we quantified the levels of fecal coliforms, E. coli, and enterococci present in collected wildfowl scats, observing a considerable variation in each of these. Of the 50 fresh fecal scats analyzed, more than half contained no detectable enterococci, with the remainder having between 10² and 10⁶ enterococci per g of fecal material (mean, 9.9 x 10⁴ g^–1). Fecal coliforms and E. coli were also only detected in less than half of the scats examined, with mean levels (1.1 x 10⁶ g^–1) at least 10-fold higher than those of enterococci. While the initial aim was to collect scats immediately after defecation, this was not often possible, so visibly fresh wildfowl scats were collected, usually first thing in the morning. It is probable that some die-off of both E. coli and enterococci occurred between excretion and analysis in the laboratory, so we may be underestimating the levels of bacterial indicators in the scats. The high solid content of the fecal samples necessitated a 10² dilution of the fecal samples so as to achieve readable plates. This resulted in a detection limit of 100 CFU/g. The use of a most probable number enrichment approach may have recovered bacterial indicators from the samples where none were isolated. This would, however, have produced clonal bacteria unsuitable for ARA. The observed prevalence is, however, consistent with previously published work where highly variable levels of enterococci and E. coli have been found in individual wild bird feces (16, 19). For example, Fallacara et al. (5) failed to isolate E. coli from a third of the feces of free-living waterfowl they tested. Previous estimates of average levels of enterococci were 10⁵ to 10⁷ g^–1 in gull feces (7, 31), 10³ to 10⁴ g^–1 in Canadian goose feces (15, 19), and 10⁵ g^–1 in whistling swan feces (15). The median levels of E. coli (5.0 x 10³ g^–1) and enterococci (2.0 x 10³ g^–1) we observed were within these ranges.

The levels of enterococci in the pond 6 discharges exceeded 10,000/100 ml on at least 25 occasions during this study. Assuming even mixing, this would correspond to at least 10¹³ enterococci in the 150 million liters discharged daily from the ponds. The daily fecal output of different wildfowl species has been estimated to range from 40 to 50 g for starlings (30) and seagulls (31) and up to 250 to 300 g for Canada geese and whistling swans (15). Even using the maximum level of enterococci observed in a scat in this study (1.5 x 10⁶) and a high daily output of 250 g of feces per bird, 10¹³ enterococci would correspond to the dedicated inputs of almost 40,000 birds. This number increases by at least 10-fold when average values are used, assumes no die-off within the ponds, and in all cases well exceeds the observed number of wildfowl on the ponds.

An unusual feature of the high levels of enterococci in pond 6 was the absence of any concurrent increase in fecal coliform levels. In wildfowl scats, fecal coliform levels were at least 10-fold higher than enterococcal levels, suggesting that if high enterococcal levels in the ponds were from wildfowl, fecal coliform levels should be even higher—this was not the case. The levels of fecal coliforms in raw sewage consistently were higher than enterococcal levels. The relative levels of the two indicators can be affected by differential decay rates, which are influenced by temperature and the environment in which they are present. While enterococci have been observed to survive longer than E. coli in an estuarine environment, (17), in oxidation ponds such as those in this study, E. coli bacteria have been found to persist longer than enterococci (25).

ARA was used to analyze enterococci from sewage, wildfowl, and pond 6. The municipal sewage contained a diverse range of enterococci, reflecting the mixed inputs of a population of more than 300,000 people. Generating a large number of profiles from the same sample resulted in very few duplicate profiles, with 482 different profiles observed among 517 isolates. Each wildfowl scat also contained a diverse range of antibiotic profiles. Of the scats where 20 or more isolates were analyzed, at least 16 and up to 34 different ARA profiles were recovered. Microbes may develop resistance to antibiotics under selective pressure, or they may acquire antibiotic resistance determinants without direct exposure to an antibiotic (18). Previous work has also observed antibiotic-resistant enterococci present in the fecal scats of wildfowl and other animals, with a wide range of ARA profiles observed (7, 19, 28).

The limited number of ARA profiles among the isolates from oxidation pond 6 was a surprising finding, with just 32 different profiles observed in the 440 isolates. These profiles were also very similar to one another and were not observed in any of the wildfowl scats or sewage samples. Using the dominant profile observed on 16 February 2006 as a reference, 72% of the isolates from pond 6 differed by resistance at two or fewer antibiotic concentrations and 88% differed at four concentrations or fewer. This degree of homogeneity was not observed in any wildfowl scat or the sewage samples. Finding the same profile in a single sample was surprising enough, but finding the same dominant profile in samples taken months apart was very unexpected. PFGE analysis confirmed that these isolates were genetically indistinguishable.

The first conclusion we could draw from this work was that the high levels of enterococci in pond 6 were not reflective of breakthrough in the sewage treatment process. By several statistical measures, the ARA profiles of the enterococci in pond 6 were very different from those of the sewage isolates. Based on available data, it is also unlikely that wildfowl were the source of the enterococci in pond 6. The small size of our library of wildfowl isolates does leave open the possibility of additional sources that we did not sample being similar to those at pond 6. However, all of the scats sampled contained enterococci with many different ARA profiles. To the best of our knowledge, there are no published reports of any animal feces containing enterococci with just a single ARA profile.

The WWTP ponds are fenced to prevent access by large wildlife and livestock, and since they are part of a protected bird sanctuary, extensive trapping and baiting systems are in operation to remove any threats to the birds from small mammals such as possums or stoats. In the absence of any other plausible fecal source, two remaining conclusions are that either environmental selection of a specific Enterococcus subtype has occurred or the high levels of enterococci are due to growth in the ponds. The high diversity of ARA types in the sources sampled would require very large inputs for the selective survival process to be able to produce the very high levels of clonal enterococci observed (up to 180,000 CFU/100 ml). We favor the growth of enterococci as the most likely explanation for the high levels observed, although we have no direct experimental evidence, such as mesocosm experiments, to support this hypothesis. The clonal nature of the isolates, as measured by both ARA and PFGE, suggests that it is a specific subset of enterococci that are growing rather than general growth of all enterococci. The wildfowl do defecate in the ponds, and if all enterococci are capable of growth, one would expect the same variety of ARA profiles as observed in wildfowl scats. It is also unusual that the fecal coliforms do not show evidence of growth—these generally have less fastidious growth requirements than enterococci. The genetic characteristics these E. faecalis strains may have, which enable them to grow preferentially to other strains, remain to be identified. In addition to these genetic characteristics, undoubtedly the key to the observed levels of enterococci, is the interaction of these enterococci with other physical, chemical, or biological factors in the ponds. Samples with elevated levels of enterococci tended to be warmer and more turbid and had higher levels of some nutrients. Whether these are significant factors in the growth of enterococci or just incidental to it, remains to be determined. The role of sediments in the persistence of enterococci and other bacteria is well documented (2, 4), with levels of up to 10⁶ CFU 100 g^–1 observed in intertidal sediments (6). Increased persistence, and possibly growth, of enterococci has been observed in association with zooplankton (22) and algal mats of Cladophora (21). Both the inhibition of protozoan predators in sediments (4) and addition of lake plankton (3) to sand can allow the growth of fecal coliforms. To our knowledge, growth of enterococci to the levels observed in the Christchurch oxidation pond has only been reported in laboratory experiments, where 4-log increases in enterococci were observed in association with the green alga Cladophora (27). The oxidation ponds appear to present a relatively controlled environment for the future study of this phenomenon.

On the basis of the evidence presented in this paper, the Christchurch City Council successfully applied to have enterococcal limits removed from their discharge conditions. The council now monitors for E. coli and, although it is not required, continues to collect data on enterococci, elevated levels of which continue to be observed sporadically. Provided these enterococci are not in themselves pathogenic, that pathogens have not also grown, and that these enterococci are not fecally associated, the presence of elevated levels of enterococci will not be associated with an elevated health risk.

Future research will attempt to characterize a broad spectrum of conditions that result in this apparent growth or environmental selection of enterococci in the absence of growth of fecal coliforms and address whether this growth can be predicted or controlled. Laboratory mesocosm experiments will be useful in understanding this phenomenon, although, as noted above, it may well involve the interaction of a number of biological organisms and factors. We also hope to identify more rapid methods that can be used to differentiate these environmental enterococci from enterococci which actually do indicate a fecal contamination source.

 
We are most grateful to Bruce Wiggins, James Madison University, for sharing protocols and advice on the use of ARA. The critical comments on the manuscript from Sandro Wolf, Megan Devane, and the four anonymous reviewers were most useful. We acknowledge the cooperation of Christchurch City Council staff in carrying out the work described here.

We acknowledge the Foundation for Research Science and Technology (FRST) for funding this research.

 
* Corresponding author. Mailing address: Institute of Environmental Science & Research Limited, Christchurch Science Centre, 27 Creyke Road, P.O. Box 29-181, Christchurch, New Zealand. Phone: 64-3-351 0044. Fax: 64-3-351 0010. E-mail: brent.gilpin{at}esr.cri.nz

Published ahead of print on 3 October 2008.

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1
1. Barrow, G. I., and R. K. A. Feltham (ed.). 1993. Cowan and Steel's manual for the identification of medical bacteria, 3rd ed. Cambridge University Press, Cambridge, United Kingdom.
2
2. Byappanahalli, M., M. Fowler, D. Shively, and R. Whitman. 2003. Ubiquity and persistence of Escherichia coli in a midwestern coastal stream. Appl. Environ. Microbiol. 69:4549-4555.[Abstract/Free Full Text]
3
3. Byappanahalli, M. N., R. L. Whitman, D. A. Shively, W. T. Ting, C. C. Tseng, and M. B. Nevers. 2006. Seasonal persistence and population characteristics of Escherichia coli and enterococci in deep backshore sand of two freshwater beaches. J. Water Health 4:313-320.[Medline]
4
4. Davies, C. M., J. A. Long, M. Donald, and N. J. Ashbolt. 1995. Survival of fecal microorganisms in marine and freshwater sediments. Appl. Environ. Microbiol. 61:1888-1896.[Abstract]
5
5. Fallacara, D. M., C. M. Monahan, T. Y. Morishita, and R. F. Wack. 2001. Fecal shedding and antimicrobial susceptibility of selected bacterial pathogens and a survey of intestinal parasites in free-living waterfowl. Avian Dis. 45:128-135.[CrossRef][Medline]
6
6. Ferguson, D. M., D. F. Moore, M. A. Getrich, and M. H. Zhowandai. 2005. Enumeration and speciation of enterococci found in marine and intertidal sediments and coastal water in southern California. J. Appl. Microbiol. 99:598-608.[CrossRef][Medline]
7
7. Fogarty, L. R., S. K. Haack, M. J. Wolcott, and R. L. Whitman. 2003. Abundance and characteristics of the recreational water quality indicator bacteria Escherichia coli and enterococci in gull faeces. J. Appl. Microbiol. 94:865-878.[CrossRef][Medline]
8
8. Gilpin, B. J., J. E. Gregor, and M. G. Savill. 2002. Identification of the source of faecal pollution in contaminated rivers. Water Sci. Technol. 46:9-15.[Medline]
9
9. Gilpin, B. J., F. Nourozi, D. Saunders, P. Grounds, P. Scholes, and M. Savill. 2003. The use of chemical and molecular microbial indicators for faecal source identification. Water Sci. Technol. 47(3):39-43.[Medline]
10
10. Graves, L. M., and B. Swaminathan. 2001. PulseNet standardized protocol for subtyping Listeria monocytogenes by macrorestriction and pulsed-field gel electrophoresis. Int. J. Food Microbiol. 65:55-62.[CrossRef][Medline]
11
11. Gregor, J., N. Garrett, B. Gilpin, C. Randall, and D. Saunders. 2002. Use of classification and regression tree (CART) analysis with chemical faecal indicators to determine sources of contamination. N. Z. J. Mar. Freshw. Res. 36:387-398.
12
12. Grundmann, H., S. Hori, and G. Tanner. 2001. Determining confidence intervals when measuring genetic diversity and the discriminatory abilities of typing methods for microorganisms. J. Clin. Microbiol. 39:4190-4192.[Abstract/Free Full Text]
13
13. Harwood, V. J., J. Whitlock, and V. Withington. 2000. Classification of antibiotic resistance patterns of indicator bacteria by discriminant analysis: use in predicting the source of fecal contamination in subtropical waters. Appl. Environ. Microbiol. 66:3698-3704.[Abstract/Free Full Text]
14
14. Hunter, S. B., P. Vauterin, M. A. Lambert-Fair, M. S. Van Duyne, K. Kubota, L. Graves, D. Wrigley, T. Barrett, and E. Ribot. 2005. Establishment of a universal size standard strain for use with the PulseNet standardized pulsed-field gel electrophoresis protocols: converting the national databases to the new size standard. J. Clin. Microbiol. 43:1045-1050.[Abstract/Free Full Text]
15
15. Hussong, D., J. M. Damare, R. J. Limpert, W. J. Sladen, R. M. Weiner, and R. R. Colwell. 1979. Microbial impact of Canada geese (Branta canadensis) and whistling swans (Cygnus columbianus columbianus) on aquatic ecosystems. Appl. Environ. Microbiol. 37:14-20.[Abstract/Free Full Text]
16
16. Kuntz, R., P. Hartel, K. Rodgers, and W. Segars. 2004. Presence of Enterococcus faecalis in broiler litter and wild bird feces for bacterial source tracking. Water Res. 38:3551-3557.[Medline]
17
17. Lessard, E. J., and J. M. Sieburth. 1983. Survival of natural sewage populations of enteric bacteria in diffusion and batch chambers in the marine environment. Appl. Environ. Microbiol. 45:950-959.[Abstract/Free Full Text]
18
18. McGinnis, S., and T. L. Madden. 2004. BLAST: at the core of a powerful and diverse set of sequence analysis tools. Nucleic Acids Res. 32:W20-W25.[Abstract/Free Full Text]
19
19. Middleton, J. H., and A. Ambrose. 2005. Enumeration and antibiotic resistance patterns of fecal indicator organisms isolated from migratory Canada geese (Branta canadensis). J. Wildl. Dis. 41:334-341.[Medline]
20
20. Naser, S. M., F. L. Thompson, B. Hoste, D. Gevers, P. Dawyndt, M. Vancanneyt, and J. Swings. 2005. Application of multilocus sequence analysis (MLSA) for rapid identification of Enterococcus species based on rpoA and pheS genes. Microbiology 151:2141-2150.[Abstract/Free Full Text]
21
21. Olapade, O. A., M. M. Depas, E. T. Jensen, and S. L. McLellan. 2006. Microbial communities and fecal indicator bacteria associated with Cladophora mats on beach sites along Lake Michigan shores. Appl. Environ. Microbiol. 72:1932-1938.[Abstract/Free Full Text]
22
22. Signoretto, C., G. Burlacchini, M. Lleo Mdel, C. Pruzzo, M. Zampini, L. Pane, G. Franzini, and P. Canepari. 2004. Adhesion of Enterococcus faecalis in the nonculturable state to plankton is the main mechanism responsible for persistence of this bacterium in both lake and seawater. Appl. Environ. Microbiol. 70:6892-6896.[Abstract/Free Full Text]
23
23. Simpson, E. H. 1949. Measurement of diversity. Nature 163:688.[CrossRef]
24
24. Sinton, L. W., R. K. Finlay, and D. J. Hannah. 1998. Distinguishing human from animal faecal contamination in water: a review. N. Z. J. Mar. Freshw. Res. 32:323-348.
25
25. Sinton, L. W., C. H. Hall, P. A. Lynch, and R. J. Davies-Colley. 2002. Sunlight inactivation of fecal indicator bacteria and bacteriophages from waste stabilization pond effluent in fresh and saline waters. Appl. Environ. Microbiol. 68:1122-1131.[Abstract/Free Full Text]
26
26. Whitlock, J. E., D. T. Jones, and V. J. Harwood. 2002. Identification of the sources of fecal coliforms in an urban watershed using antibiotic resistance analysis. Water Res. 36:4273-4282.[Medline]
27
27. Whitman, R. L., D. A. Shively, H. Pawlik, M. B. Nevers, and M. N. Byappanahalli. 2003. Occurrence of Escherichia coli and enterococci in Cladophora (Chlorophyta) in nearshore water and beach sand of Lake Michigan. Appl. Environ. Microbiol. 69:4714-4719.[Abstract/Free Full Text]
28
28. Wiggins, B. A. 1996. Discriminant analysis of antibiotic resistance patterns in fecal streptococci, a method to differentiate human and animal sources of fecal pollution in natural waters. Appl. Environ. Microbiol. 62:3997-4002.[Abstract]
29
29. Wiggins, B. A., P. W. Cash, W. S. Creamer, S. E. Dart, P. P. Garcia, T. M. Gerecke, J. Han, B. L. Henry, K. B. Hoover, E. L. Johnson, K. C. Jones, J. G. McCarthy, J. A. McDonough, S. A. Mercer, M. J. Noto, H. Park, M. S. Phillips, S. M. Purner, B. M. Smith, E. N. Stevens, and A. K. Varner. 2003. Use of antibiotic resistance analysis for representativeness testing of multiwatershed libraries. Appl. Environ. Microbiol. 69:3399-3405.[Abstract/Free Full Text]
30
30. Wither, A., M. Rehfisch, and G. Austin. 2005. The impact of bird populations on the microbiological quality of bathing waters. Water Sci. Technol. 51:199-207.[Medline]
31
31. Wood, A. J., and T. J. Trust. 1972. Some qualitative and quantitative aspects of the intestinal microflora of the glaucous-winged gull (Larus glaucescens). Can. J. Microbiol. 18:1577-1583.[Medline]

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Applied and Environmental Microbiology, December 2008, p. 7204-7210, Vol. 74, No. 23
0099-2240/08/$08.00+0     doi:10.1128/AEM.00341-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

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