East Fork Lake (8.7 km²; maximum depth 33.4 m) and Delaware Lake (3.9 km²; maximum depth 9.1 m) are human-made reservoirs constructed by the United States(US) Army Corps of Engineers for flood control, water supply, recreation, and wildlife management in the Olentangy Watershed. Madison Lake is a small (0.4 km²) and shallow (maximum depth 1.8 m) lake for agricultural land use in the Darby Plains of Madison County (Lee et al., 2014). The lakes are often influenced by non-point pollution sources from the surrounding watersheds, including croplands, livestock pasture, forestland, and residential areas. The water quality can be also impacted by human fecal contamination through recreational activities during summer as the lakes are used for recreational purposes, such as swimming beach, picnic areas, and shelterhouses for many visitors (http://ohiodnr.gov).

A total of 21 water samples were collected from three recreational freshwaters in Ohio: 9 samples from East Fork Lake (39° 01′ 96.2″ N; 84° 13′ 41.2″ W), 6 samples from Delaware Lake (40° 37′ 14.6″ N; 83° 05′ 87.2″ W), and 5 samples Madison Lake (39° 86′ 97.5″ N; 83° 37′ 43.8″ W; Figure 1). From the surface (~30 cm) near the center of each lake (1-m water depth), a single daily sample (>2 L) was collected in the afternoon (between 12:00 and 18:00) on weekends from June to August 2009 using two autoclaved 1 L bottles (Nalgene, Rochester, NY). Temperature, pH, specific conductivity, and dissolved oxygen level were measured with the YSI 600XL multiprobe data sonde (Yellow Springs Instruments, Yellow Springs, OH) in triplicate. Turbidity was measured with the Hach 2100P portable turbidimeter [(US Environmental Protection Agency (USEPA), 1983)]. The sampling was carried in accordance with the recommendations of the Ohio Department of Health Sampling Guideline.

After transportation to the laboratory on ice, a 200 mL aliquot of each water sample was filtered through a 47-mm-diameter, 0.22-μm-pore-sized mixed cellulose ester membrane filter (Millipore, Bedford, MA) to collect bacteria (Yannarell and Triplett, 2005). The membrane filter was transferred into a 50 mL sterile tube and then stored at −80°C until further analysis. To fecal indicator levels, E. coli densities were determined in triplicate within 6 h after transportation to the laboratory according to USEPA Method 1603 (USEPA, 2002). Precipitation data were obtained local weather stations of the National Weather Service (https://www.ncdc.noaa.gov/data-access). Water inflow data were obtained from the Louisville District of the US Army Corps of Engineers (http://www.lrl.usace.army.mil/) and the National Water Information System of the US Geological Survey (http://waterdata.usgs.gov/oh/nwis/rt).

Bacterial DNA was extracted from the cellulose membrane as described previously (Wolcott et al., 2009). Briefly, the membrane filter was cut into two pieces using sterile scalpel blades and each piece was transferred into a sterile microcentrifuge tube containing 500 μL RLT buffer with β-mercaptoethanol (Qiagen, Valencia, CA). For complete bacterial lysis, a bead-beating method was performed using a Qiagen TissueLyser at 30 Hz for 5 min with a sterile 5 mm steel bead (Qiagen) and 500 μL sterile 0.1 mm glass beads (Scientific Industries, Inc., NY). After centrifugation at 14,000 rpm for 30 s, 100 μL supernatant aliquots from each tube were transferred into a new sterile microcentrifuge tube. Thus a total of 200 μL supernatatnts were prepared in the same tube and subjected to DNA extraction using a Qiagen Stool Kit (Qiagen). Extracted DNA was eluted from the DNA spin column into 30 μL DNA/RNA-free water and diluted to a final concentration of 20 ng/μL using a NanoDrop spectrophotometer (Nyxor Biotech, Paris, France). Positive control for the DNA extraction was performed by spiking Bacteroides fragilis ATCC 25285^T into 200 ml of sterile phosphate buffered saline (PBS; 0.14 M NaCl, 2.7 mM KCl, 10.1 mM Na₂HPO₄, 1.8 mM KH₂PO₄ [pH 7.6]) as described previously (Lee et al., 2014). For a negative control, 200 mL of sterile phosphate buffered saline without spiking was filtered through a membrane filter and the DNA extraction was carried out from the membrane filter, resulting in no false-positive signals.

Bacterial tag-encoded pyrosequencing was performed at the Research and Testing Laboratory (Lubbock, TX) using the Roche 454/FLX system according to the manufacturer's instructions (Roche, Indianapolis, IN). The 16S universal eubacterial primers Gray28F (5′-GAGTTTGATCNTGGCTCAG-3′) and Gray519R (5′-GTNTTACNGCGGCKGCTG-3′) were used to amplify up to the 520 bp V1-V3 region of the 16S rRNA gene as described previously (Dowd et al., 2008). The resultant amplicons were analyzed through a pipeline of the Research and Testing Laboratory (http://www.researchandtesting.com/docs/Data_Analysis_Methodology.pdf). Briefly, noisy sequence-reads were initially removed by screening for low quality (<25 Phred scores), short length (<250 bps), and long homopolymers (>6 bp). Singleton sequences and ambiguous bases in the regions of the primer and barcode were removed. Prefix de-replication, clustering, and centroid sequence generation were performed using the USEARCH algorithm (Edgar, 2010). Further non-flow based denoising was performed using the Acacia software (ver. 1.52) (Bragg et al., 2012) to correct errors. Analysis of the sequencing data was performed using bioinformatics programs, as implemented in the QIIME software package (ver. 1.4.0) (Caporaso et al., 2010b). Sequences were aligned against the Greengenes core set database using the PyNAST option (Caporaso et al., 2010a), and chimeric sequences were removed from the aligned sequences using the ChimeraSlayer option (Haas et al., 2011). Quality-checked sequences were taxonomically assigned based on the Ribosomal Database Project (RDP) naïve Bayesian rRNA Classifier v2.2 (Wang et al., 2007) and clustered into operational taxonomic units (OTUs) at 97% sequence similarity as a cut-off value for the species assignment using the UCLUST option (Edgar, 2010). For a more detailed taxonomical classification, the sequences assigned to 13 tentative Cyanobacteria genera (GpI to GPXIII) were reclassified based on the National Center for Biotechnology Information (NCBI) database. All pyrosequencing reads obtained in this study were submitted to the NCBI Sequence Read Archive (SRA) under the study accession number SAMN02087719–SAMN02087739.

The observed OTUs, estimated OTUs (Chao1 and ACE), and alpha diversity indices (Shannon and Simpson) were calculated with normalized sequence reads using the QIIME software (Caporaso et al., 2010b). The maximum number of OTUs and coverage percentages were calculated from rarefaction curves using the non-linear model procedure (PROC NLIN) of the SAS software (V9.1; SAS Institute, Cary, NC) as described previously (Kim et al., 2011). Principal coordinates analysis (PCoA) was conducted using the weighted UniFrac distance matrix of a phylogenetic tree constructed with representative OTUs (Lozupone and Knight, 2005), which was compared with the hierarchic clustering constructed by the hclust function in the R software package using average linkage algorithms (UPGMA) (R Development Core Team, 2009).

To assess potential associations among environmental, fecal indicator, and pyrosequencing data sets, Spearman correlation and canonical correlation analyses (CCA) were conducted using the R package (ver. 2.15.1) (González et al., 2008). The data for the microbiological terms, including fecal indicator levels and pyrosequencing reads, were log (base 10)-transformed. To compare non-normally distributed data of environmental parameters among the three lakes, the Kruskal-Wallis test was performed using the R package, where significant differences and tendencies were set to P < 0.05 and 0.1, respectively.

For the evaluation of toxic cyanobacteria, one of microcystin biosynthesis gene cluster, mcyA gene, was measured in triplicate by a SYBR qPCR assay using a primer set (forward M1rF: 5′-AGCGGTAGTCATTGCATCGG-3′, reverse M1rR: 5′-GCCCTTTTTCTGAAGTCGCC-3′) (Yoshida et al., 2007). For the quantification of the mcyA genetic marker, the DNA of a Microcystis aeruginosa NIES-843 culture (as a positive control; kindly provided by Dr. Wilhelm, University of Tennessee) was extracted by the xanthogenate nucleic acid isolation method (Tillett and Neilan, 2000). The mcyA qPCR assay was carried out in a total volume of 20 μL qPCR mixture containing a 2μL of M. aeruginosa DNA template, 10 μL of SYBR universal PCR master mix (Applied Biosystems, Foster City, CA), and a 200 nM primer set. Thermal cycling was conducted for an initial 95°C for 10 min, followed by 40 cycles of denaturation at 95°C for 30 s, and annealing and extension at 62°C for 3 min using the ABI 48-well StepOne™ Real Time System (Applied Biosystems, Foster City, CA).

To check specificity of qPCR amplicons in the assay, a melting point analysis was carried out by lowering the temperature to 50°C, and then raising it by 1.0°C/s until it reached 95°C. A mixture of all PCR reagents without DNA/RNA templates was used as a negative control, resulting in no false-positive signals. For the standard curve, the PCR products of mcyA gene were cloned using the pGEM-T Vector System (Promega Co., Madison, WI), followed by the blue-white screening and sub-culturing according to the manufacturer's instruction. The plasmid DNAs of sub-cultured clones were extracted using QIAprep Spin Miniprep Kit (Qiagen, Valencia, CA). The concentration of DNA was measured with NanoDrop spectrophotometer and each 10-fold serial dilution was then prepared using linearized plasmid DNA, followed by each qPCR assay to construct each standard curve with plotting C_T values vs. log₁₀ values of the gene copy number, where a limit of quantification (LOQ) was defined as the lowest gene copy number within the linear range of quantification (Lee et al., 2015).

Environmental parameters of water samples are summarized by lake (Table 1). In general, dissolved oxygen, turbidity, water inflow, and precipitation had normal variations without significant differences among the three lakes. However, higher pH and lower specific conductivity were found in East Fork Lake compared to the other two lakes (P < 0.001). The temperature of East Fork Lake tended to be higher than those of the two other lakes (P < 0.1).

For microbial water quality parameters, higher geometric means of E. coli were found in Madison Lake compared to East Fork Lake or Delaware Lake (P < 0.05). E. coli densities of 18 out of the 21 samples remained below the single-sample advisory limit in freshwater beaches (235 CFU/100 mL) defined by USEPA advisory. Three high E. coli densities above the advisory limit were observed from two East Fork Lake samples (Aug 1 and 2) and one Madison Lake sample on July 13 (Supplementary Figure 1A). Levels of mcyA genetic marker ranged from undetected to 4.3 × 10³ gene copies/100 mL; positive detections were observed from two East Fork Lake samples (June 21 and June 27), two Delaware Lake samples (July 20 and July 21), and one Madison Lake sample (June 24) (Supplementary Figure 1B).

The level of mcyA genetic marker was calculated using a standard curve. The standard curve showed a good linear range of quantification from 4.4 × 10⁰ to 4.4 × 10⁸ gene copies/reaction (R² = 0.9963). The slope, y-intercept, and amplification efficiency of the standard curve were −3.5334, 35.728, and 91.8%, respectively. The limit of quantification was determined as 4.4 × 10⁰ gene copies/reaction (Supplementary Figure 2).

A total of 108,566 quality-checked sequences (average length 430 bp; average Phred score 34) were retrieved from 21 total samples from three lakes. To normalize read number variation among samples, raw sequences in the range of 3396–7141 per sample were normalized into the minimum number (3396) by selecting proportional percentages from each dataset with the subsample function in the QIIME. The re-sampled dataset was used to estimate the bacterial diversity and abundance. At the species level, defined by 97% sequence similarity, the numbers of OTUs ranged from ~300 to 700; where the coverage ratio of observed OTUs to estimated OTUs ranged from 66 to 88%. The Shannon and Simpson indices of samples ranged from 2.641 to 4.564 and 0.053 to 0.228 at 97% sequence similarity, respectively (Table 2).

Total, shared, and unique OTUs from the three lakes are illustrated in a weighted Venn diagram (Figure 2). East Fork Lake had the most unique bacteria (1534 OTUs), followed by Madison Lake (1303 OTUs) and Delaware Lake (540 OTUs). Shared bacterial components were observed; 678 OTUs between East Fork Lake and Delaware Lake, 628 OTUs were shared between East Fork Lake and Madison Lake, and 487 OTUs between Delaware Lake and Madison Lake, respectively. Common bacteria found in all three lakes were 334 OTUs, which belonged to the genera Cyanobacteria GpIIa, Legionella, Methylocystis, Mycobacterium, and Polynucleobacter (Figure 2).

The microbial communities included four dominant phyla (Actinobacteria, Cyanobacteria, Firmicutes, and Proteobacteria) with more than 95% of the total population, as well as less abundant phyla (Acidobacteria, Bacteroidetes, Chlorobi, Chloroflexi, Deinococcus-Thermus, Fusobacteria, Gemmatimonadetes, Planctomycetes, Synergistetes, Tenericutes, Verrucomicrobia, and a candidate phylum TM7) with <5% of the total population. In general, Actinobacteria were most abundant in the three lakes (30.0% in East Fork Lake, 41.9% in Delaware Lake, and 35.2% in Madison Lake. Cyanobacteria and Firmicutes were also dominant; however, their abundances differed between East Fork Lake and the other two lakes. Cyanobacteria were more abundant in East Fork Lake (33.0%) than in the other lakes (11.6% in Delaware Lake and 13.8% in Madison Lake), while Firmicutes were less abundant in East Fork Lake (8.5%) than the other lakes (34.7% in Delaware Lake and 27.3% in Madison Lake). Proteobacteria accounted for 19.8% in East Fork Lake, 8.7% in Delaware Lake, and 17.9% in Madison Lake (Figure 3A). Temporal variations were observed in the relative abundances of some bacterial classes such as Actinobacteria, Bacilli, and Cyanobacteria; the class Bacilli had the most variation, ranging from 0.1 to 84.6% of the total population. On the other hand, Alpha-, Beta-, and Gammaproteobacteria showed less fluctuation in their abundances across all water samples, irrespective of lake, compared to the other dominant classes (Figure 3B).

A more detailed bacterial community structure is illustrated by a heat-map at the genus level (Figure 4). Briefly, dominant members (accounting for >5% of the total sequences) were: Cyanobacteria GpIIa (29.1%), Mycobacterium (13.4%), and Exiguobacterium (7.5%) in East Fork Lake; Paenisporosarcina (27.7%), Mycobacterium (18.5%), Cyanobacteria GpIIa (9.1%), Exiguobacterium (7.2%), and Arthrobacter (5.7%) in Delaware Lake; and Mycobacterium (12.6%), Exiguobacterium (12.5%), and Staphylococcus (6.9%) in Madison Lake.

In the Spearman correlation analysis of pyrosequencing reads, a significant positive correlation was found at the genus level between Cyanobacteria GpIIa and temperature in all lakes (r = 0.514, P = 0.017). Within bacterial compositions at the class level, Cyanobacteria and Bacilli had a negative relationship with each other (r = −0.526, P = 0.014). In addition to the negative relationship between Cyanobacteria GpIIa and Methylocystis in all lakes (r = −0.454, P = 0.039), Cyanobacteria GpIIa was negatively related to Arthrobacter (r = −0.590, P = 0.026) in Madison Lake and with Exiguobacterium (r = −0.895, P = 0.040) or Planomicrobium (r = −0.977, P = 0.004) in Delaware Lake (Table 3).

A clustering dendrogram showed that the bacterial communities were clustered into four groups as follows: (1) a mixed cluster included one East Fork Lake sample (July 26), two Delaware Lake samples (June 11 and 16), and two Madison Lake samples (June 24 and July 6); (2) a cluster with seven East Fork Lake samples (June 21 and August 1, 2, 8, 15, 16, and 22); (3) a mixed cluster with three Delaware Lake samples (July 13, 20, and 21) and four Madison Lake samples (June 30, July 14, July 21, and July 27); and (4) a cluster with one Madison Lake sample on July 13 (Figure 5A). In the PCoA plot, the range of PC1 mostly differed for the water samples of East Fork Lake (0.0–0.4) compared with those of Delaware Lake and Madison Lake (−0.4–0.0). The PC2-values were rather dispersed, regardless of lake (Figure 5B). Results from both the PCoA plot and the clustering dendrogram showed that the spatial difference in bacterial community structure was more apparent in East Fork Lake compared to the other lakes. This suggests that both the variety and abundances of bacterial in East Fork Lake differed from those in the other two lakes, as the weighted PCoA plot utilizes a qualitative β diversity reflecting changes in how many sequences from each lineage are present, as well as changes in which taxa are present (Lozupone and Knight, 2005).

CCA was used to examine potential relationships between bacterial community composition and the environmental and microbial parameters measured in this study (Figure 6). Among the parameters, temperature, pH, dissolved oxygen, precipitation, and the mcyA genetic marker were the major components of the first axis in the ordination plot, explaining 71.3% of total variance, whereas water inflow, turbidity, and E. coli were major components of the second axis, explaining 21.1% of total variance. Bacilli and Cyanobacteria were separated from each other on the horizontal axis, while Actinobacteria, Bacteroidia, Clostridia, and Gammaproteobacteria were scattered on the vertical axis compared to other bacterial taxa located near the origin point (Figure 6). CCA showed strong correlations of Clostridia and Bacteroidia with water inflow on July 13 at Madison Lake, while Gammaproteobacteria were associated with turbidity on August 1 at East Fork Lake. In addition, the two bacterial classes, Bacteroidia and Clostridia, showed similar directions with that of E. coli in the CCA plot. On June 11 at Delaware Lake and July 6 at Madison Lake, Bacilli appeared to be linked with precipitation, while Cyanobacteria showed a positive association with temperature, pH, and the mcyA genetic marker.

The taxonomical assignment of the pyrosequencing dataset showed that the phyla of Actinobacteria, Cyanobacteria, Firmicutes, and Proteobacteria prevailed in the three lakes. Actinobacteria were previously regarded as typical inhabitants of soil environments. However, Actinobacteria have been recently reported as autochthonous bacteria in freshwater habitats (Warnecke et al., 2004), that can degrade organic polymers such as a lignin (Ghai et al., 2014), or uptake amino acids by metabolizing N-containing aromatic polymers such as a chitin (Salcher et al., 2010; Eckert et al., 2012). The most abundant member in the phylum Actinobacteria was the genus Mycobacterium, which has been reported to inhabit various aquatic environments by utilizing various hydrocarbons (Vaerewijck et al., 2005). Actinobacteria was also reported to contain complete sets of genes to uptake phosphorus by coding enzymes for the conversion reaction from inositol phosphates into acetyl-CoA (Ghai et al., 2014). The co-abundance of Actinobacteria and Cyanobacteria is often observed under the eutrophic conditions with high phosphorus in water. Recently, two novel lineages in Actinobacteria, acTH1 and acTH2, were detected as dominant bacteria from both the water and sediment of shallow cyanobacteria-blooming freshwater in China, Lake Taihu (Wu Q. L. et al., 2007; Ghai et al., 2014), where all sequences belonged to the acTH2 lineage were identified as the genus Mycobacterium.

At the genus level, Synechococcus and Cyanobium were dominant components of Cyanobacteria in East Fork Lake and Delaware Lake, while the genus Planktothrix prevailed in Madison Lake. Considering that East Fork Lake and Delaware Lake are more oligotrophic than Madison Lake (Marion et al., 2010), this contrasting result is not surprising. The nutrient status of a lake can be a critical factor in the community structure shift of Cyanobacteria. The high occurrences of Synechococcus and Cyanobium in East Fork Lake and Delaware Lake were consistent with those in other oligotrophic freshwater lakes (Postius and Ernst, 1999; Callieri and Stockner, 2002). A study of Cyanobacteria in Lake Erie reported that both Synechococcus and Cyanobium were not found two decades ago, but became predominant genera due to a shift from a eutrophic to oligo-mesotrophic status (Ouellette et al., 2006). In contrast, Planktothrix has been reported as a bacterial indicator of eutrophic or hypertrophic lakes (Lira et al., 2011).

The class Bacilli exhibited large fluctuations in the phylum Firmicutes among the three lakes, raising the question of a potential link between its ecological niche and selective environmental conditions. For example, the genus Exiguobacterium was the most dominant group in the class Bacilli and was more frequently detected in Delaware Lake and Madison Lake than in East Fork Lake (Figure 4). It was reported that Bacilli were the most active bacteria in synthesizing de novo DNA for the spore germination during the early stages of re-wetting events in dried river sediments. This suggests that Bacilli could become primary microbial colonizers, and then passively release planktonic bacteria into the freshwater in their vegetative form (Fazi et al., 2008). In addition, a comparative genomic study reported that Exiguobacterium could exist in wide temperature ranges as two distinct forms, psychrophilic and thermophilic (Vishnivetskaya et al., 2009). Delaware Lake has a wide inundation area due to its large watershed, with water level changes of up to 6 m, while Madison Lake is shallow with a maximum depth of <2 m. On the other hand, the second deepest lake in Ohio, East Fork Lake, has a steep and rocky shoreline, falling off rapidly into deep water with a maximum depth of 34 m (Ohio Department of Natural Resources, http://ohiodnr.gov/). Considering these ecological and environmental conditions, the Exiguobacterium are speculated to survive and grow in the wide temperature and moisture ranges in the watershed of Delaware Lake and the sediments of Madison Lake.

In addition, Betaproteobacteria are known as some of the most abundant bacteria among the phylum Proteobacteria in typical freshwater systems (Van der Gucht et al., 2005). However, Betaproteobacteria have been reported to decrease during a cyanobacterial bloom events (Bagatini et al., 2014). The changes of pH affected by cyanobacterial growth could have an influence on the bacterial community structure of Betaproteobacteria. Among all genera in the class Betaproteobacteria, Polynucleobacter were detected in nearly all of the water samples (95%, 20/21) and P. acidiphobus was the most dominant species in the genus Polynucleobacter (Supplementary Table 1). Previous studies reported that P. acidiphobus is abundant at high pH (Wu X. et al., 2007), and not found in acidic freshwaters (Hahn et al., 2011). Thus, the distinct occurrence of P. acidiphobus suggested that the differentiation of the Betaproteobacteria community was likely influenced by high pH conditions (about 9) due to the growth of cyanobacteria.

In East Fork Lake, temperature was an important environmental factor influencing the bacterial community structure, especially for Synechococcus, which was the most representative genus of Cyanobacteria GpIIa. This is in accordance with another study reporting that temperature is the most determinative environmental factor for the population of Synechococcus (Lindström et al., 2005).

The negative correlation between Cyanobacteria and Bacilli was attributed to relationships between their dominant component genera. The genus Methylocystis is a group of Gram-negative aerobic type II methanotrophic bacteria that utilize methane as their sole source of carbon and energy, which favors environments such as lake sediments containing relatively high levels of methane and low levels of dissolved oxygen (Auman et al., 2000). The genus Arthrobacter is regarded as a group of zymogenous bacteria inhabiting soil and rhizosphere, with fluctuating abundance in response to changing soil conditions (Cacciari and Lippi, 1987). Exiguobacterium has wide distributions in soil and aquatic environments, such as permafrost/contemporary surface soil (Vishnivetskaya and Kathariou, 2005), and freshwater/seawater (Rodrigues et al., 2009). Planomicrobium is often isolated from halophilic environments such as fermented seafood, marine mud, and coastal sediment (Jung et al., 2009). The reasons for the negative correlations between Cyanobacteria and Bacilli remain unknown. However, these four psychrophilic genera of Bacilli seem to prefer cold temperature conditions (Wagner, 2008), while Cyanobacteria seem to proliferate exponentially under higher temperature conditions. In this regard, a stratified sampling with shorter intervals is needed to verify the connections between Cyanobacteria and Bacilli.

Even though harmful algal blooms were not observed in the three lakes during the study period, Cyanobacteria GI were frequently found in East Fork and Madison lakes from the pyrosequencing analysis. Most of the Cyanobacteria GI sequences were assigned to the genus Anabaena. Interestingly, the mcyA genetic marker shared the same direction with the Cyanobacteria GI cluster at the genus level in the CCA plot (data not shown). Cyanobacteria IV and XI were rarely found in water samples from East Fork Lake, which were assigned to Leptolyngbya sp. and M. aeruginosa. In Madison Lake, Planktothrix was most abundant in the cyanobacterial community, which is one of potential toxin-producing cyanobacteria (Christiansen et al., 2003).

The clustering result of the pyrosequencing dataset was interesting in Madison Lake. The family Enterobacteriaceae was significantly higher in Madison Lake (8.9%) than in the other two lakes (0.4–1.5%). In the Enterobacteriaceae, Escherichia had the greatest occurrence on July 13 in Madison Lake. The genera Bacteroides and Enterococcus were also found on the same day, where ~230 sequences (4.1% of the total bacteria) were assigned to the genus Enterococcus. Although the genera Bacteroides and Enterococcus can be derived from environmental sources, they were identified as B. fragilis and E. faecalis, which can be frequently observed in animal and human fecal flora (Simpson et al., 2002). Even so, the precise time of a fecal contamination event is unclear due to the possibility of persistent DNAs from antecedent contaminations. However, it is likely that the bacterial community structure in Madison Lake on July 13 was affected by external bacterial loading via water inflows containing fecal contamination (Figure 6).

The relative abundance of particular taxonomic groups changed significantly between water samples. Some genera of bacteria showed significantly higher abundance in the three lakes during the period of this study (e.g., Cyanobacteria, Mycobacterium, Methylocystis, and Legionella), calling for further investigation regarding whether they can be listed as candidates for opportunistic and emerging pathogens threatening public health. There were numerous unidentified OTUs in the phyla Actinobacteria and Proteobacteria. Despite no evidence for the presence of pathogenic Mycobacterium spp. (e.g., M. tuberculosis), many OTUs assigned to Mycobacterium spp. were dominant components in the water samples, including M. moriokaense, M. wolinskyi, M. brisbanense, M. arupense, and M. senegalense. Recently, these environmental or non-tuberculous mycobacteria have been known to cause opportunistic infections in humans (Primm et al., 2004).

Moreover, more than 10% of all of the Proteobacteria sequences were assigned to unclassified Legionellaceae in this study, implying a certain presence of unknown Legionellaceae in the recreational freshwaters. Many Legionella species can exist in aquatic environments and they have become important agent relevant to public health (Fields, 1996). For example, more than 18 species of Legionella were reported to be related to patients with pneumonia (Bangsborg, 1997). To clarify the potential health risks in recreational freshwaters, further isolation and characterization of unidentified Mycobacterium and Legionella are warranted.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.