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Decay of infectious adenovirus and coliphages in freshwater habitats is differentially affected by ambient sunlight and the presence of indigenous protozoa communities

Virology Journal volume 17, Article number: 1 (2020) Cite this article

Abstract

Background

Sanitary quality of recreational waters worldwide is assessed using fecal indicator bacteria (FIB), such as Escherichia coli and enterococci. However, fate and transport characteristics of FIB in aquatic habitats can differ from those of viral pathogens which have been identified as main etiologic agents of recreational waterborne illness. Coliphages (bacteriophages infecting E. coli) are an attractive alternative to FIB because of their many morphological and structural similarities to viral pathogens.

Methods

In this in situ field study, we used a submersible aquatic mesocosm to compare decay characteristics of somatic and F+ coliphages to those of infectious human adenovirus 2 in a freshwater lake. In addition, we also evaluated the effect of ambient sunlight (and associated UV irradiation) and indigenous protozoan communities on decay of somatic and F+ coliphage, as well as infectious adenovirus.

Results

Our results show that decay of coliphages and adenovirus was similar (p = 0.0794), indicating that both of these bacteriophage groups are adequate surrogates for decay of human adenoviruses. Overall, after 8 days the greatest log10 reductions were observed when viruses were exposed to a combination of biotic and abiotic factors (2.92 ± 0.39, 4.48 ± 0.38, 3.40 ± 0.19 for somatic coliphages, F+ coliphages and adenovirus, respectively). Both, indigenous protozoa and ambient sunlight, were important contributors to decay of all three viruses, although the magnitude of that effect differed over time and across viral targets.

Conclusions

While all viruses studied decayed significantly faster (p < 0.0001) when exposed to ambient sunlight, somatic coliphages were particularly susceptible to sunlight irradiation suggesting a potentially different mechanism of UV damage compared to F+ coliphages and adenoviruses. Presence of indigenous protozoan communities was also a significant contributor (p value range: 0.0016 to < 0.0001) to decay of coliphages and adenovirus suggesting that this rarely studied biotic factor is an important driver of viral reductions in freshwater aquatic habitats.

Introduction

Bacteriophages have a long history of use as model organisms in the realm of molecular biology such as the investigation of the transfer of genes, mechanisms of gene repression and activation, and various gene therapy applications [1]. While bacteriophages have been the subject of research efforts for many years [2, 3], there has been a renewed interest in recent years for practical applications in both public and environmental health arenas. In addition to a growing interest in using bacteriophages as tools to combat antibiotic resistant bacteria [4,5,6,7], there has been a recent effort to develop recreational water quality criteria for Escherichia coli infecting bacteriophages (i.e. somatic and F+ coliphages) [8].

Coliphages have been used routinely in many monitoring programs (e.g. ground water, aquaculture practices, water reuse, biosolids) [9,10,11] and rationale for their inclusion in recreational water quality assessment [8, 12] is that their persistence in aquatic habitats can more closely resemble that of viral pathogens because of many morphological and structural similarities [13]. While sanitary quality of recreational waters is routinely assessed through enumeration of fecal indicator bacteria (FIB, such as E. coli and enterococci), recent reports identifying viral pathogens as leading causes of recreational waterborne diseases outbreaks [14,15,16,17] combined with known differences in fate and transport between FIB and viruses [18,19,20,21,22,23] highlights the need to evaluate suitability of viral indicators to predict pathogen decay in environmental waters.

Although removal of FIB and viruses through primary and secondary wastewater treatment processes is similar [24,25,26,27], viruses are reported to display a greater resilience to wastewater disinfection practices compared to FIB [28,29,30,31], allowing them to enter recreational waters through treated wastewater discharge. In contrast, others have shown that reduction of coliphages and viral pathogens through wastewater treatment processes is comparable [18, 32] suggesting that they are similarly affected by exposure to different physical and chemical stressors (e.g. chlorination, UV, peracetic acid, etc). While frequent co-occurrence of coliphages and viral pathogens in environmental waters [33,34,35,36,37,38,39], often in the absence of FIB, implies a similar response to various biotic and abiotic environmental stressors, field studies examining this are rare.

Some studies investigating drivers of decay for both coliphage and viral pathogens have suggested that their response to certain environmental stressors is similar. For example, both groups tend to persist longer at lower temperatures [40,41,42,43] and in freshwater as compared to marine waters [44,45,46]. On the other hand, while decay of infectious coliphages is accelerated when exposed to ambient and simulated sunlight [44, 47,48,49,50,51], the response of pathogenic viruses is more ambiguous [50,51,52] and possibly influenced by laboratory measurement strategies (infectious viruses enumerated on mammalian cell cultures versus molecular approaches such as qPCR enumerating viral nucleic acids) [52, 53]. Even less is known about the potential effect of biotic stressors, such as protozoan predation, on decay of both coliphages and viral pathogens. Greater decay in the presence of indigenous microbiota has been demonstrated for FIB and some bacterial pathogens [54,55,56,57], but analogous information is needed for viruses.

Factors impacting viral persistence in natural systems are difficult to simulate, necessitating an experimental design that closely mimics natural conditions. To address these research gaps, we employed a submersible aquatic mesocosm (SAM) to study decay of coliphages (somatic and F+) and infectious adenoviruses in a freshwater lake under in situ conditions. We also investigated the effect of indigenous protozoan communities and ambient sunlight to better understand the biotic and abiotic factors impacting the decay of viruses in natural aquatic environment.

Materials and methods

Experimental design

Ambient water (~ 15 L) was collected from William H. Harsha Lake (Batavia, OH: 39.0252°N, − 84.1303° W). Immediately after collection, 50% of the sample was passed through a 0.80 μm filter to remove indigenous protozoa. Filtration of water to remove protozoa is a common method and more effective than other techniques such as chemical treatments [58,59,60,61,62]. To minimize any changes in microbial populations, filtered and unfiltered water was stored in dark at 4 °C until the beginning of the experiment (< 48 h). In order to closely mimic ambient conditions by in situ incubation (at William H. Harsha Lake), a SAM was used to conduct the study. The SAM was constructed as previously described [54, 63,64,65,66] and samples were contained using regenerated cellulose dialysis bags (75 mm flat width, 13–14 kD pore size molecular weight cut-off, Spectrum Labs, Rancho Dominguez, CA). The first day of the experiment, both filtered and unfiltered ambient water was spiked with somatic and F+ coliphages and adenovirus and stirred for 15 min to ensure proper distribution of the spikes within the sample. Measured portions of either spiked filtered or spiked unfiltered ambient water (200 mL) were used to fill each dialysis bag. One half of the dialysis bags containing each water type was attached at the top portion (approximately 2–5 cm below the water surface for the light exposure treatment), while the other half was placed at the bottom portion (approximately 25–30 cm below the water surface underneath the heavy-duty black plastic tarp for shaded treatment). For the study, four different treatments were as follows: A: exposure to ambient sunlight and indigenous microbiota including protozoa (top level, unfiltered water), B: exposure to indigenous microbiota including protozoa, (bottom level, unfiltered water), C: exposure to ambient sunlight only (top level, filtered water) and D: exposure to neither variable (bottom level, filtered water). During each sampling event, triplicate dialysis bags for each treatment were processed for the enumeration of somatic and F+ coliphages, as well as infectious adenovirus (as described below). Concentrations of all viruses were obtained immediately after the inoculum preparation (day 0) and after one and eight days of exposure. Two additional time points (days 3 and 5) were processed for both coliphage types.

Bacteriophage enumeration

Somatic and F+ coliphage were enumerated using double agar layer (DAL) procedure, as previously described [67]. If necessary, decimal dilution series were prepared using 1X phosphate buffered saline solution (PBS: 0.0425 g/L KH2PO4 and 0.4055 g/L of MgCl2; pH 7.2 Sigma Aldrich, St. Louis, MO). Briefly, 1 mL of sample was added to 5 mL of “molten” top tryptic soy agar (TSA) layer (0.7% agar) containing 0.1% of appropriate antibiotic stock solution (100 μg/ mL nalidixic acid for somatic and 15 μg/ mL streptomycin/ampicillin for F+ coliphage) (Fisher Scientific, Waltham, MA), followed by addition of 200 μl of appropriate E. coli host (CN-13 ATCC#700609 [somatic] of Famp ATCC#700891 [F+], American Type Culture Collection, Manassas, VA) in mid-log growth phase. The soft agar overlay mixture was mixed and poured on bottom agar TSA plates (1.5% agar and containing 0.1% of appropriate antibiotic stock solution). Plates were incubated at 37 °C for 16–18 h. The following day characteristic plaque forming units (PFU) for each coliphage type were enumerated and data were expressed as PFU per 1 ml. Method blank (sample substituted with 1X PBS) and media sterility negative controls were performed on each day of the experiment. For the duration of the study, no plaques were observed on any of the negative controls indicating absence of contamination.

Adenovirus enumeration

Human lung cells (A549, ATCC® CCL-185) were propagated in Dulbecco’s Minimum Essential Medium (DMEM high glucose with HEPES, Greiner, Monroe, NC) supplemented with 10% fetal calf serum (Fisher Scientific) and 1% sodium pyruvate (Fisher Scientific) under 5% CO2 atmosphere and at 37 °C. Test cultures of A549 cells were planted and grown to 90% confluency for 4 days in 25 cm2 filter capped flasks at 37 °C using a maintenance medium (as described above) except: 1) the addition of antibiotic-antimycotic solution (1% v/v, Fisher Scientific) and 2) reduced fetal calf serum amount of 2% v/v. Prior to inoculation with samples, test cultures were washed with 10 mL of Earle’s Balanced Salt Solution per flask (EBSS, Fisher Scientific) supplemented with 1% antibiotic-antimycotic solution. Decimal dilution series of samples were created using 1X PBS and five replicate flasks per dilution were utilized. In addition, ten negative control flasks (containing 10 mL of 1X PBS instead of the sample) were ran with each sample batch. Following inoculation, flasks were placed on a rocker for 90 min to allow for viral attachment/infection to occur. Flasks were then supplemented with 10 mL of maintenance medium and incubated at 37 °C for 3 weeks [68]. During the incubation time, flasks were examined weekly for the formation of cytopathic effects (CPE). Concentrations of adenovirus were estimated using EPA’s Most Probable Number (MPN) calculator Version 2.0 (https://cfpub.epa.gov/si/si_public_record_report.cfm?Lab=NERL&dirEntryId=309398). Resulted are reported as MPN per 1 mL.

Virus spike preparation

Primary treated wastewater was collected from a local wastewater treatment plant and used as a source of somatic and F+ coliphages. Briefly, 10 mL of wastewater was syringe filtered (0.22 μm pore size) and added to 100 mL of mid-log culture of appropriate E. coli host. The inoculated host cultures were incubated at 37 °C for 16-18 h, followed by centrifugation (3800 x g, 15 min) and syringe filtration (0.22 μm pore size). The resulting coliphage stocks were titered using DAL as described above and stored in dark at 4 °C until the beginning of the experiment (~ 24 h).

Human adenovirus 2 (ATCC® VR-846) was obtained from ATCC and propagated in A549 cells to generate higher titers. Briefly, A549 cells were infected with adenovirus as described above for the samples. Following the development of CPE (typically in < a week), cells underwent three freeze-thaw cycles, followed by centrifugation at 2500 x g for 30 min to pellet cellular debris. The supernatant was syringe filtered (0.22 μm pore size), titered (as described above for cell culture samples) and stored in dark at 4 °C until the beginning of the experiment.

Visible light and temperature measurements

For the duration of the study, hourly light intensity (lum/ft2) and temperature (°C) measurements were recorded at both upper and lower SAM levels using HOBO® UA 002–08 data loggers (Onset Computer Corporation, Bourne, MA). The temperature at the top level (16.67 ± 1.18 °C) was slightly higher (paired t-test, p = 0.0394) compared to the bottom level 16.59 ± 0.88 °C), but the light intensity was considerably greater (paired t-test, p < 0.0001) at the top (54.34 ± 146.73 lum/ft2) compared to the bottom level (9.47 ± 19.15 lum/ft2).

Data analysis

All concentration data were log10 transformed prior to data analyses. Log10 reductions were calculated by subtracting concentrations obtained on day “n” (where “n” represents days 1, 3, 5 or 8) from concentration at the beginning of the experiment (day 0). GraphPad Prism version 7.01 (GraphPad Software, La Jolla, CA) was used to conduct a two-way analysis of variance (ANOVA with interactions) with Tukey’s multiple comparison test to evaluate the effects of two factors (indigenous microbiota including protozoa and sunlight) on decay. This software was also used to conduct the paired t-tests, one-way ANOVA and Pearson product momentum correlation to assess significant differences in light temperature measurements, across different virus measurements and to identify potential correlations trends in decay patterns, respectively.

Results

Decay characteristics in freshwater environment

Overall, average log10 reduction on days one and eight for all treatments was greatest for adenovirus (1.48 ± 0.99), followed by F+ (0.79 ± 1.53) and somatic (0.61 ± 1.21) coliphages, although these differences were not statistically significant (p = 0.0794). After 8 days, exposure to sunlight and indigenous microbiota (Treatment A) resulted in the greatest decay for all three organisms (log10 reductions of 2.92 ± 0.39, 4.48 ± 0.38, 3.41 ± 0.19 for somatic coliphages, F+ coliphages and adenovirus, respectively) (Table 1, Figs 1, 2 and 3). Exposure to sunlight only (Treatment C) resulted in log10 reductions of 2.31 ± 0.20, 1.17 ± 0.01 and 1.54 ± 0.24 for somatic coliphages, F+ coliphages and adenovirus respectively, while shaded treatments (Treatments B and D) typically yielded the least decay (log10 range: 0.05–1.11) (Table 1, Figs 1, 2 and 3). Decay pattern of all viruses was strongly correlated (r2 range: 0.754–0.881, p value range: 0.0002 - < 0.0001), although it was the most noticeable for F+ coliphage and adenoviruses (r2 = 0.881, p < 0.0001).

Table 1 Log10 reduction values for somatic coliphage, F+ coliphage and adenovirus. Treatments: A (exposure to sunlight and indigenous microbiota including protozoa), B (exposure to only indigenous microbiota including protozoa), C (exposure to sunlight only), D (exposure to neither)
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Fig. 1
figure 1

Effect of treatment variables on changes in somatic coliphage concentrations over time. Error bars represent standard deviation. Treatments: A (exposure to sunlight and indigenous microbiota including protozoa), B (exposure to only indigenous microbiota including protozoa), C (exposure to sunlight only), D (exposure to neither)

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Fig. 2
figure 2

Effect of treatment variables on changes in F+ coliphage concentrations over time. Error bars represent standard deviation. Treatments: A (exposure to sunlight and indigenous microbiota including protozoa), B (exposure to only indigenous microbiota including protozoa), C (exposure to sunlight only), D (exposure to neither)

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Fig. 3
figure 3

Effect of treatment variables on changes in adenovirus concentrations over time. Error bars represent standard deviation. Treatments: A (exposure to sunlight and indigenous microbiota including protozoa), B (exposure to only indigenous microbiota including protozoa), C (exposure to sunlight only), D (exposure to neither)

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Effect of experimental variables on decay of somatic coliphages

After 1 day of exposure, neither variable (ambient sunlight and protozoan grazing) had a significant effect on decay, and log10 reductions for all treatments were negligible. Over the course of the next 48 h (day three), unfiltered treatments containing indigenous protozoa exhibited greater decay (log10 reduction values: 1.17 ± 0.04 and 1.40 ± 0.12; (Treatments A and B, respectively) compared to filtered treatments (< 1 log10 reduction; Treatments C and D) (Table 1, Fig. 2). While presence of protozoa was the only significant variable affecting decay at day three (Table 2) and contributing ~ 86% to variation in the data set, the interaction between variables was also significant (p = 0.0026) indicating that the effect of protozoa was dependent on sunlight exposure (Table 2). Forty-eight hours later (day five), effects of protozoa began to diminish (Fig. 1, Table 1), and sunlight irradiation became the dominant factor affecting the decay (log10 reduction values: 1.34 ± 0.17 and 1.66 ± 0.19 and for Treatments A and C, respectively) and contributing ~ 94% to the observed variation in decay (Table 2). At day eight, which was the final time point, solar irradiation continued to be the dominant factor (~ 95% contribution to the observed variability in the dataset) and was the only influential variable (Table 2) causing the log10 reduction values of 2.92 ± 0.39 and 2.32 ± 0.20 for Treatments A and C, respectively (Table 1, Fig. 1).

Table 2 Two-way ANOVA with Tukey’s post-hoc test of treatment effects during each time point. Statistically significant values are bolded
Full size table

Effect of experimental variables on decay of F+ coliphages

Similar to somatic coliphages, decay of F+ coliphages was minimal within the first 24 h of exposure and neither variable had significant effect on decay (Tables 1 and 2, Fig. 2). At day three, 48 h later, exposure to indigenous microbiota had a significant effect on decay contributing ~ 50% to the observed variations in the data set with minimal interactions (Table 2).

The greatest decay occurred in Treatment A (exposure to sunlight and biota; log10 1.62 ± 0.22), followed by Treatment B (exposure to biota only; log10 1.14 ± 0.17) and finally Treatments C and D (exposure to sun only and exposure to neither variable; < 1.00 log10 reduction each) (Table 1, Fig. 2). Presence of protozoa continued to significantly affect the decay of F+ coliphages at day five, but its contribution to the variability was less (~ 18%). Exposure to sunlight became a dominant significant variable on day five, contributing nearly 50% to the observed variation in the dataset (Table 2). Overall, the greatest decay occurred for the treatment containing both, indigenous protozoa and sunlight (log10 reduction: 3.16 ± 0.11), indicating that the effect of variables was co-dependent (Table 1). During the final time point (day eight), the effect of both variables (as well as their interaction) continued to be statistically significant and their contribution to the decay remained similar to that at day five (p = 0.0001; Table 2). Decay continued to be the most pronounced when F+ coliphage were exposed to both variables (log10 reduction Treatment A: 4.48 ± 0.38), followed by exposure to sunlight only (log10 reduction Treatment C: 1.43 ± 0.10) while the decay in the remaining two treatments was negligible (Table 1, Fig. 2).

Effect of experimental variables on decay of infectious adenovirus 2

Although decay data for adenovirus is limited, similar to both coliphages, neither variable had a significant effect on decay within the first 24 h of exposure (Table 2, Fig. 3). Over the course of 8 days, both sunlight and indigenous microbiota were significant factors contributing to the decay of adenovirus (Table 2, Fig. 3). Sunlight was more important variable contributing nearly 50% to the observed variations in the data set, followed by interactions between the variables (~ 24%) and indigenous biota (~ 19%) (Table 2). In sunlight treatments, adenovirus reduction in presence of protozoa (Treatment A) was approximately 2 log10 greater compared to the reductions in their absence (Treatment C) (3.41 ± 0.19 vs 1.54 ± 0.24) (Table 1, Fig. 3), whereas the reduction in dark treatments was ~ 1 log10.

Discussion

Recent reports indicate that the majority of recreational waterborne illnesses are caused by viral pathogens [14,15,16,17]. As a result, routine monitoring of recreational waters with FIB may not adequately represent viral pathogen presence due [69], at least in part, to different decay trends between these two groups [18]. Coliphages are an attractive alternative because they have similar morphological characteristics to those of many pathogenic viruses suggesting they can better mimic their survival compared to FIB [70, 71]. Earlier studies reported that somatic and F+ coliphages were adequate surrogates for fate and transport of poliovirus [72] and noroviruses [73], respectively. In this in situ field study we used a SAM to compare the decay characteristics of somatic and F+ coliphages to that of infectious adenovirus and to evaluate the effect of ambient sunlight and indigenous protozoan communities on their decay.

The effect of ambient sunlight (and associated UV-A and UV-B radiation) on decay of various indicators and pathogens is likely one of the most commonly studied abiotic environmental factors [74]. Briefly, the damage caused by ambient sunlight can be classified into two categories, direct and indirect based on the mode of action [75]. Direct damage is caused by UV-B and it results in a formation of pyrimidine dimers, while UV-A causes indirect, photooxidative damage which can be exogenous or endogenous depending on the location of free radicals and reactive oxygen species [75]. Earlier studies noted that the decay of infectious adenoviruses exposed to natural and simulated sunlight [50, 76, 77] was typically greater than their corresponding qPCR signal [52, 53, 78] in both marine and freshwaters. Similar findings were observed for infectious somatic and F+ coliphages [44, 50, 66, 76]. We also noted a strong influence of ambient sunlight on decay of infectious coliphages and adenovirus 2, especially after 5 days of exposure, although it is worth noting that we used a singular, laboratory propagated strain of adenovirus and that indigenous, environmental strains may exhibit greater resilience [79,80,81]. Furthermore, the effect of ambient sunlight was more pronounced for somatic coliphages, compared to F+ coliphages and adenoviruses. This is consistent with previous studies [50, 66, 76, 82] reporting a greater susceptibility of somatic coliphages to sunlight compared to other viral groups. While exposure to both UV-A and UV-B spectrum is detrimental, earlier studies investigating the mechanism of sunlight action, suggest that indirect, photooxidative damage may be the primary mechanism for adenovirus and F+ coliphages [44, 51, 76], while direct damage caused by UV-B is the dominant mechanism for somatic coliphages [44, 76]. However, additional controlled, laboratory based mechanistic studies are needed to confirm that the greater susceptibility of somatic coliphages, as compared to F+ coliphages and adenoviruses, to sunlight is due to differential decay modes of action.

Ciliates and heterotrophic nanoflagellates are effective grazers in the water column [83] and an important part of microbial food webs in many different aquatic habitats [84]. The abundance of these two groups in oligo-mesotrophic waters, such as William H. Harsha Lake is typically estimated to be between 102 and 104 cells per mL [85, 86]. While the effects of protozoan predation have been demonstrated for FIB and other bacteria in field studies [54,55,56, 66], the role biotic interactions play in decay of viruses is rarely explored. Laboratory feeding experiments demonstrated uptake of various adenoviruses (serotypes 2, 11 and 41) by ciliate Euplotes octocarinatus [87] and a free-living amoeba, Acanthamoeba castellanii [88], as well as adsorption of adenovirus 2 on the surface of wild ciliates isolated from active sludges of a wastewater treatment plant [87]. However, direct immunofluorescent antibody techniques were used to detect adenoviruses inside and on the surface of the protozoan cells [87, 88] and it is unclear whether the viruses were infectious. Laboratory decay studies conducted in the dark and in the absence of indigenous microbiota (autoclaved ground and river water) noted extended persistence of infectious adenovirus 2 and 41 [89, 90], but the faster decay of infectious poliovirus type 1 was noted in the presence of indigenous microbiota (compared to autoclaved controls) [91], suggesting that indigenous microbiota play an important role in the decay of infectious viruses.

Like adenovirus laboratory feeding experiments, a recent report demonstrated macropinocytosis and digestion of T4 coliphage in food vacuoles of ciliate Tetrahymena thermophila [92], suggesting that active virophagy by protozoans in environmental waters may be an important mechanism for viral attenuation. Similarly, ingestion by suspension feeding heterotrophic flagellates Thaumatomonas coloniensis and Salpingoeca spp. (rather than adsorption) was demonstrated for MS2 coliphage in groundwater [93]. Furthermore, some studies suggest that MS2 coliphage may be a source of nutrients for predatory protozoa [93, 94], further supporting the notion that predation may be an important biotic factor influencing viral decay. The limited number of field studies suggest that the removal of enterophages (bacteriophages infecting Enterococcus faecalis) [95] and F+ coliphages [96] is greater in unamended lake and river waters compared to the filtered and/or autoclaved controls, but decay of latter group appears to be subgroup specific [96]. However, a marine water in situ study showed a minimal effect of indigenous microbiota on decay of somatic and F+ coliphages, as well as GB-124 bacteriophage infecting Bacteroides fragilis [66], suggesting that the effect of protozoan communities on viral decay may be influenced by water type (fresh versus marine).

We observed a significant reduction of infectious adenovirus 2 and both coliphage groups (although it was more pronounced for the F+ compared to somatic coliphage) in the presence of indigenous protozoa and under the influence of ambient sunlight. This was especially pronounced after 3 to 5 days of exposure to indigenous protozoan communities, a trend that is consistent with the time required for freshwater protozoan communities to adjust to the influx of prey organisms [97,98,99]. This finding suggests that indigenous protozoa likely plays an important role in the decay of infectious viruses (indicators and pathogens alike), especially in freshwater habitats and in conjunction with ambient sunlight, although the magnitude of that effect is influenced by the time point and the viral target. Future studies are needed to clarify the nature of ecological interactions between protozoans and viruses and to better characterize the interplay between sunlight irradiation and impact of indigenous protozoa on viral decay.

Conclusions

In summary, our results indicate that both somatic and F+ coliphages decay at similar rates to infectious adenoviruses in a freshwater aquatic habitat. This finding implies that their persistence in environmental waters could be similar and that coliphages may be suitable surrogates for adenovirus decay in these systems. Furthermore, while we show that the exposure to ambient sunlight plays an important role in viral decay, its effect was especially pronounced with somatic coliphages, suggesting that the mechanism of action may differ among the viruses studied. Lastly, our data suggests that protozoans play an important role in the decay of somatic and F+ coliphages and infectious adenoviruses in aquatic environments. While controlled laboratory-based studies can provide important insights into the effect of environmental factors on decay, additional field studies closely mimicking natural conditions are warranted to better characterize the interactions between indigenous protozoan communities and infectious viral pathogens and indicators.

Availability of data and materials

Data can be found on EPA ScienceHub website (https://catalog.data.gov/harvest/epa-sciencehub).

Abbreviations

ANOVA:

Analysis of variance

ATCC:

American type culture collection

CPE:

Cytopathic effects

DAL:

Double agar layer

DMEM:

Dulbecco’s minimum essential medium

EBSS:

Earle’s balanced salt solution

FIB:

Fecal indicator bacteria

MPN:

Most probable number

PBS:

Phosphate buffered saline

SAM:

Submersible aquatic mesocosm

TSA:

Tryptic soy agar

UV:

Ultraviolet

References

  1. Flint SJ, Enquist LW, Racaniello VR, Skalka AM. Principles of Virology, 3rd edition, Volume I: Molecular Biology. Washington, D.C.: ASM Press; 2009.

    Google Scholar 

  2. Casjens SR, Hendrix RW. Bacteriophage lambda: Early pioneer and still relevant. Virology. 2015;479-480:310–30.

    CAS  PubMed  Article  Google Scholar 

  3. Lobocka M, Szybalski WT. Advances in virus research, Bacteriophages PART B. New York: Elsevier; 2012.

    Google Scholar 

  4. Wang M, Xiong W, Liu P, Xie X, Zeng J, Sun Y, Zeng Z. Metagenomic Insights Into the Contribution of Phages to Antibiotic Resistance in Water Samples Related to Swine Feedlot Wastewater Treatment. Front Microbiol. 2018;9:2474.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  5. Wang M, Liu P, Zhou Q, Tao W, Sun Y, Zeng Z. Estimating the contribution of bacteriophage to the dissemination of antibiotic resistance genes in pig feces. Environ Pollut. 2018;238:291–8.

    CAS  PubMed  Article  Google Scholar 

  6. Ross J, Topp E. Abundance of Antibiotic Resistance Genes in Bacteriophage following Soil Fertilization with Dairy Manure or Municipal Biosolids, and Evidence for Potential Transduction. Appl Environ Microbiol. 2015;81:7905–13.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  7. Lekunberri I, Villagrasa M, Balcazar JL, Borrego CM. Contribution of bacteriophage and plasmid DNA to the mobilization of antibiotic resistance genes in a river receiving treated wastewater discharges. Sci Total Environ. 2017;601-602:206–9.

    CAS  PubMed  Article  Google Scholar 

  8. United States Environmental Protection Agency. Review of coliphages as possible indicators of fecal contamination for ambient water quality. Washington, D.C; 2015.

  9. North Carolina Environmental Quality. North Carolina Adm. Code 15A NCAC 2U Reclaimed Water. Raleigh: North Carolina Department of Environment and Natural Resources; 2011.

    Google Scholar 

  10. Queensland Government Environmental Protection Agency. Queensland Water Recycling Guidelines. Brisbane; 2005.

  11. United States Environmental Protection Agency. National Primary Drinking Water Regulations: Groundwater rule. Final Rule; 40 CFR Parts 9, 141 and 142, Federal Register, vol. 71. Washington, D.C; 2006. p. 216.

  12. United States Environmental Protection Agency. 2016 Coliphage Experts Workshop: Discussion Topics and Findings. Washington, D.C; 2016.

  13. King AMQ, Adams MJ, Carstens EB, Lefkowitz WJ. Virus Taxonomy: Classification and Nomenclature of Viruses. Ninth Report of the International Committee on Taxonomy of Viruses. London: Elsevier Academic Press; 2011.

    Google Scholar 

  14. Sinclair RG, Jones EL, Gerba CP. Viruses in recreational water-borne disease outbreaks: a review. J Appl Microbiol. 2009;107:1769–80.

    CAS  PubMed  Article  Google Scholar 

  15. Graciaa DS, Cope JR, Roberts VA, Cikesh BL, Kahler AM, Vigar M, Hilborn ED, Wade TJ, Backer LC, Montgomery SP, Evan Secor W, Hill VR, Beach MJ, Fullerton KE, Yoder JS, Hlavsa MC. Outbreaks Associated with Untreated Recreational Water - United States, 2000-2014. Am J Transplant. 2018;18:2083–7.

    Article  Google Scholar 

  16. Guzman-Herrador B, Carlander A, Ethelberg S, Freiesleben de Blasio B, Kuusi M, Lund V, Lofdahl M, MacDonald E, Nichols G, Schonning C, Sudre B, Tronnberg L, Vold L, Semenza JC, Nygard K. Waterborne outbreaks in the Nordic countries, 1998 to 2012. Euro Surveill. 2015;20:24.

    PubMed  Article  Google Scholar 

  17. Hlavsa MC, Roberts VA, Kahler AM, Hilborn ED, Mecher TR, Beach MJ, Wade TJ, Yoder JS, Centers for Disease C, Prevention. Outbreaks of Illness Associated with Recreational Water--United States, 2011-2012. MMWR Morb Mortal Wkly Rep. 2015;64:668–72.

    PubMed  PubMed Central  Google Scholar 

  18. McMinn BR, Ashbolt NJ, Korajkic A. Bacteriophages as indicators of faecal pollution and enteric virus removal. Lett Appl Microbiol. 2017;65:11–26.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  19. Ahmed W, Gyawali P, Sidhu JP, Toze S. Relative inactivation of faecal indicator bacteria and sewage markers in freshwater and seawater microcosms. Lett Appl Microbiol. 2014;59:348–54.

    CAS  PubMed  Article  Google Scholar 

  20. Characklis GW, Dilts MJ, Simmons OD 3rd, Likirdopulos CA, Krometis LA, Sobsey MD. Microbial partitioning to settleable particles in stormwater. Water Res. 2005;39:1773–82.

    CAS  PubMed  Article  Google Scholar 

  21. Cho KH, Pachepsky YA, Oliver DM, Muirhead RW, Park Y, Quilliam RS, Shelton DR. Modeling fate and transport of fecally-derived microorganisms at the watershed scale: State of the science and future opportunities. Water Res. 2016;100:38–56.

    CAS  PubMed  Article  Google Scholar 

  22. Ferguson C, Husman AMD, Altavilla N, Deere D, Ashbolt N. Fate and transport of surface water pathogens in watersheds. Crit Rev Environ Sci Technol. 2003;33:299–361.

    Article  Google Scholar 

  23. Gantzer C, Gillerman L, Kuznetsov M, Oron G. Adsorption and survival of faecal coliforms, somatic coliphages and F-specific RNA phages in soil irrigated with wastewater. Water Sci Technol. 2001;43:117–24.

    CAS  PubMed  Article  Google Scholar 

  24. Lucena F, Duran AE, Moron A, Calderon E, Campos C, Gantzer C, Skraber S, Jofre J. Reduction of bacterial indicators and bacteriophages infecting faecal bacteria in primary and secondary wastewater treatments. J Appl Microbiol. 2004;97:1069–76.

    CAS  PubMed  Article  Google Scholar 

  25. Ottoson J, Norstrom A, Dalhammar G. Removal of micro-organisms in a small-scale hydroponics wastewater treatment system. Lett Appl Microbiol. 2005;40:443–7.

    CAS  PubMed  Article  Google Scholar 

  26. Aw TG, Gin KYH. Environmental surveillance and molecular characterization of human enteric viruses in tropical urban wastewaters. J Appl Microbiol. 2010;109:716–30.

    CAS  PubMed  Google Scholar 

  27. Lodder WJ, de Roda Husman AM. Presence of noroviruses and other enteric viruses in sewage and surface waters in The Netherlands. Appl Environ Microbiol. 2005;71:1453–61.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  28. Tree JA, Adams MR, Lees DN. Chlorination of indicator bacteria and viruses in primary sewage effluent. Appl Environ Microbiol. 2003;69:2038–43.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  29. Duran AE, Muniesa M, Moce-Llivina L, Campos C, Jofre J, Lucena F. Usefulness of different groups of bacteriophages as model micro-organisms for evaluating chlorination. J Appl Microbiol. 2003;95:29–37.

    CAS  PubMed  Article  Google Scholar 

  30. Gehr R, Wagner M, Veerasubramanian P, Payment P. Disinfection efficiency of peracetic acid, UV and ozone after enhanced primary treatment of municipal wastewater. Water Res. 2003;37:4573–86.

    CAS  PubMed  Article  Google Scholar 

  31. Harwood VJ, Levine AD, Scott TM, Chivukula V, Lukasik J, Farrah SR, Rose JB. Validity of the indicator organism paradigm for pathogen reduction in reclaimed water and public health protection. Appl Environ Microbiol. 2005;71:3163–70.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  32. Tree JA, Adams MR, Lees DN. Disinfection of feline calicivirus (a surrogate for Norovirus) in wastewaters. J Appl Microbiol. 2005;98:155–62.

    CAS  PubMed  Article  Google Scholar 

  33. Boehm AB, Yamahara KM, Love DC, Peterson BM, McNeill K, Nelson KL. Covariation and photoinactivation of traditional and novel indicator organisms and human viruses at a sewage-impacted marine beach. Environ Sci Technol. 2009;43:8046–52.

    CAS  PubMed  Article  Google Scholar 

  34. Marie V, Lin J. Viruses in the environment - presence and diversity of bacteriophage and enteric virus populations in the Umhlangane River, Durban, South Africa. J Water Health. 2017;15:966–81.

    PubMed  Article  Google Scholar 

  35. Verani M, Federigi I, Donzelli G, Cioni L, Carducci A. Human adenoviruses as waterborne index pathogens and their use for Quantitative Microbial Risk Assessment. Sci Total Environ. 2019;651:1469–75.

    CAS  PubMed  Article  Google Scholar 

  36. Espinosa AC, Arias CF, Sanchez-Colon S, Mazari-Hiriart M. Comparative study of enteric viruses, coliphages and indicator bacteria for evaluating water quality in a tropical high-altitude system. Environ Health. 2009;8:49.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  37. Jiang S, Noble R, Chu W. Human adenoviruses and coliphages in urban runoff-impacted coastal waters of Southern California. Appl Environ Microbiol. 2001;67:179–84.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  38. Moce-Llivina L, Lucena F, Jofre J. Enteroviruses and bacteriophages in bathing waters. Appl Environ Microbiol. 2005;71:6838–44.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  39. Ballester NA, Fontaine JH, Margolin AB. Occurrence and correlations between coliphages and anthropogenic viruses in the Massachusetts Bay using enrichment and ICC-nPCR. J Water Health. 2005;3:59–68.

    CAS  PubMed  Article  Google Scholar 

  40. Meschke JS, Sobsey MD. Comparative reduction of Norwalk virus, poliovirus type 1, F+ RNA coliphage MS2 and Escherichia coli in miniature soil columns. Water Sci Technol. 2003;47:85–90.

    CAS  PubMed  Article  Google Scholar 

  41. Fong TT, Lipp EK. Enteric viruses of humans and animals in aquatic environments: health risks, detection, and potential water quality assessment tools. Microbiol Mol Biol Rev. 2005;69:357–71.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  42. Allwood PB, Malik YS, Maherchandani S, Hedberg CW, Goyal SM. Effect of temperature on the survival of F-specific RNA coliphage, feline calicivirus, and Escherichia coli in chlorinated water. Int J Environ Res Public Health. 2005;2:442–6.

    CAS  PubMed  Article  Google Scholar 

  43. Romero OC, Straub AP, Kohn T, Nguyen TH. Role of temperature and Suwannee River natural organic matter on inactivation kinetics of rotavirus and bacteriophage MS2 by solar irradiation. Environ Sci Technol. 2011;45:10385–93.

    CAS  PubMed  Article  Google Scholar 

  44. Sinton LW, Hall CH, Lynch PA, Davies-Colley RJ. Sunlight inactivation of fecal indicator bacteria and bacteriophages from waste stabilization pond effluent in fresh and saline waters. Appl Environ Microbiol. 2002;68:1122–31.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  45. Jeanneau L, Solecki O, Wery N, Jarde E, Gourmelon M, Communal PY, Jadas-Hecart A, Caprais MP, Gruau G, Pourcher AM. Relative decay of fecal indicator bacteria and human-associated markers: a microcosm study simulating wastewater input into seawater and freshwater. Environ Sci Technol. 2012;46:2375–82.

    CAS  PubMed  Article  Google Scholar 

  46. Hurst CJ, Gerba CP. Stability of simian rotavirus in fresh and estuarine water. Appl Environ Microbiol. 1980;39:1–5.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  47. Noble RT, Lee IM, Schiff KC. Inactivation of indicator micro-organisms from various sources of faecal contamination in seawater and freshwater. J Appl Microbiol. 2004;96:464–72.

    CAS  PubMed  Article  Google Scholar 

  48. Sinton LW, Finlay RK, Lynch PA. Sunlight inactivation of fecal bacteriophages and bacteria in sewage-polluted seawater. Appl Environ Microbiol. 1999;65:3605–13.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  49. Sokolova E, Astrom J, Pettersson TJ, Bergstedt O, Hermansson M. Decay of Bacteroidales genetic markers in relation to traditional fecal indicators for water quality modeling of drinking water sources. Environ Sci Technol. 2012;46:892–900.

    CAS  PubMed  Article  Google Scholar 

  50. Love DC, Silverman A, Nelson KL. Human virus and bacteriophage inactivation in clear water by simulated sunlight compared to bacteriophage inactivation at a southern California beach. Environ Sci Technol. 2010;44:6965–70.

    CAS  PubMed  Article  Google Scholar 

  51. Silverman AI, Peterson BM, Boehm AB, McNeill K, Nelson KL. Sunlight inactivation of human viruses and bacteriophages in coastal waters containing natural photosensitizers. Environ Sci Technol. 2013;47:1870–8.

    CAS  PubMed  Article  Google Scholar 

  52. Bae S, Wuertz S. Survival of host-associated bacteroidales cells and their relationship with Enterococcus spp., Campylobacter jejuni, Salmonella enterica serovar Typhimurium, and adenovirus in freshwater microcosms as measured by propidium monoazide-quantitative PCR. Appl Environ Microbiol. 2012;78:922–32.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  53. Ogorzaly L, Bertrand I, Paris M, Maul A, Gantzer C. Occurrence, survival, and persistence of human adenoviruses and F-specific RNA phages in raw groundwater. Appl Environ Microbiol. 2010;76:8019–25.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  54. Korajkic A, McMinn BR, Shanks OC, Sivaganesan M, Fout GS, Ashbolt NJ. Biotic interactions and sunlight affect persistence of fecal indicator bacteria and microbial source tracking genetic markers in the upper Mississippi river. Appl Environ Microbiol. 2014;80:3952–61.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  55. Korajkic A, Wanjugi P, Harwood VJ. Indigenous microbiota and habitat influence Escherichia coli survival more than sunlight in simulated aquatic environments. Appl Environ Microbiol. 2013;79:5329–37.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  56. Wanjugi P, Harwood VJ. The influence of predation and competition on the survival of commensal and pathogenic fecal bacteria in aquatic habitats. Environ Microbiol. 2013;15:517–26.

    PubMed  Article  Google Scholar 

  57. Wanjugi P, Harwood VJ. Protozoan predation is differentially affected by motility of enteric pathogens in water vs. sediments. Microb Ecol. 2014;68:751–60.

    PubMed  Article  Google Scholar 

  58. Oemcke DJ, Hans van Leeuwen J. Ozonation of the marine dinoflagellate alga Amphidinium sp.--implications for ballast water disinfection. Water Res. 2005;39:5119–25.

    CAS  PubMed  Article  Google Scholar 

  59. Doblin MA, Dobbs FC. Setting a size-exclusion limit to remove toxic dinoflagellate cysts from ships' ballast water. Mar Pollut Bull. 2006;52:259–63.

    CAS  PubMed  Article  Google Scholar 

  60. Batani G, Perez G, de la Escalera GM, Piccini C, Fazi S. Competition and protist predation are important regulators of riverine bacterial community composition and size distribution. J Freshw Ecol. 2016;31:609–23.

    CAS  Article  Google Scholar 

  61. Worsfold NT, Warren PH, Petchey OL. Context-dependent effects of predator removal from experimental microcosm communities. Oikos. 2009;118:1319–26.

    Article  Google Scholar 

  62. Suttle CA, Chen F. Mechanisms and rates of decay of marine viruses in seawater. Appl Environ Microbiol. 1992;58:3721–9.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  63. Korajkic A, McMinn BR, Ashbolt NJ, Sivaganesan M, Harwood VJ, Shanks OC. Extended persistence of general and cattle-associated fecal indicators in marine and freshwater environment. Sci Total Environ. 2019;650:1292–302.

    CAS  PubMed  Article  Google Scholar 

  64. Korajkic A, Parfrey LW, McMinn BR, Baeza YV, VanTeuren W, Knight R, Shanks OC. Changes in bacterial and eukaryotic communities during sewage decomposition in Mississippi river water. Water Res. 2015;69:30–9.

    CAS  PubMed  Article  Google Scholar 

  65. Korajkic A, McMinn BR, Harwood VJ, Shanks OC, Fout GS, Ashbolt NJ. Differential decay of enterococci and Escherichia coli originating from two fecal pollution sources. Appl Environ Microbiol. 2013;79:2488–92.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  66. Wanjugi P, Sivaganesan M, Korajkic A, Kelty CA, McMinn B, Ulrich R, Harwood VJ, Shanks OC. Differential decomposition of bacterial and viral fecal indicators in common human pollution types. Water Res. 2016;105:591–601.

    CAS  PubMed  Article  PubMed Central  Google Scholar 

  67. Adams MH. Bacteriophages. New York: Interscience Publishers; 1959.

    Google Scholar 

  68. Cashdollar JL, Huff E, Ryu H, Grimm AC. The influence of incubation time on adenovirus quantitation in A549 cells by most probable number. J Virol Methods. 2016;237:200–3.

    CAS  PubMed  Article  Google Scholar 

  69. Korajkic A, McMinn BR, Harwood VJ. Relationships between Microbial Indicators and Pathogens in Recreational Water Settings. Int J Environ Res Public Health. 2018;15:12.

    CAS  PubMed Central  Article  Google Scholar 

  70. Payment P, Plante R, Cejka P. Removal of indicator bacteria, human enteric viruses, Giardia cysts, and Cryptosporidium oocysts at a large wastewater primary treatment facility. Can J Microbiol. 2001;47:188–93.

    CAS  PubMed  Article  Google Scholar 

  71. Payment P, Locas A. Pathogens in water: value and limits of correlation with microbial indicators. Ground Water. 2011;49:4–11.

    CAS  PubMed  Article  Google Scholar 

  72. Skraber S, Gassilloud B, Schwartzbrod L, Gantzer C. Survival of infectious Poliovirus-1 in river water compared to the persistence of somatic coliphages, thermotolerant coliforms and Poliovirus-1 genome. Water Res. 2004;38:2927–33.

    CAS  PubMed  Article  Google Scholar 

  73. Bae J, Schwab KJ. Evaluation of murine norovirus, feline calicivirus, poliovirus, and MS2 as surrogates for human norovirus in a model of viral persistence in surface water and groundwater. Appl Environ Microbiol. 2008;74:477–84.

    CAS  PubMed  Article  Google Scholar 

  74. Korajkic A, Wanjugi P, Brooks L, Cao Y, Harwood VJ. Persistence and Decay of Fecal Microbiota in Aquatic Habitats. Microbiol Mol Biol Rev. 2019;83:e00005–19.

    PubMed  Article  PubMed Central  Google Scholar 

  75. Nelson KL, Boehm AB, Davies-Colley RJ, Dodd MC, Kohn T, Linden KG, Liu YY, Maraccini PA, McNeill K, Mitch WA, Nguyen TH, Parker KM, Rodriguez RA, Sassoubre LM, Silverman AI, Wigginton KR, Zepp RG. Sunlight-mediated inactivation of health-relevant microorganisms in water: a review of mechanisms and modeling approaches. Environ Sci Processes Impacts. 2018;20:1089–122.

    CAS  Article  Google Scholar 

  76. Mattle MJ, Vione D, Kohn T. Conceptual model and experimental framework to determine the contributions of direct and indirect photoreactions to the solar disinfection of MS2, phiX174, and adenovirus. Environ Sci Technol. 2015;49:334–42.

    CAS  PubMed  Article  Google Scholar 

  77. Elmahdy MEI, Magri ME, Garcia LA, Fongaro G, Barardi CRM. Microcosm environment models for studying the stability of adenovirus and murine norovirus in water and sediment. Int J Hyg Environ Health. 2018;221:734–41.

    CAS  PubMed  Article  Google Scholar 

  78. Liang L, Goh SGA, Gin KYH. Decay kinetics of microbial source tracking (MST) markers and human adenovirus under the effects of sunlight and salinity. Sci Total Environ. 2017;574:165–75.

    CAS  Article  Google Scholar 

  79. Ravva SV, Sarreal CZ. Persistence of F-Specific RNA Coliphages in Surface Waters from a Produce Production Region along the Central Coast of California. PLoS One. 2016;11:e0146623.

    PubMed  PubMed Central  Article  CAS  Google Scholar 

  80. Brion GM, Meschke JS, Sobsey MD. F-specific RNA coliphages: occurrence, types, and survival in natural waters. Water Res. 2002;36:2419–25.

    CAS  PubMed  Article  Google Scholar 

  81. Balleste E, Blanch AR. Persistence of Bacteroides species populations in a river as measured by molecular and culture techniques. Appl Environ Microbiol. 2010;76:7608–16.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  82. Wu J, Cao Y, Young B, Yuen Y, Jiang S, Melendez D, Griffith JF, Stewart JR. Decay of Coliphages in Sewage-Contaminated Freshwater: Uncertainty and Seasonal Effects. Environ Sci Technol. 2016;50:11593–601.

    CAS  PubMed  Article  Google Scholar 

  83. Zhang J, Ormala-Odegrip AM, Mappes J, Laakso J. Top-down effects of a lytic bacteriophage and protozoa on bacteria in aqueous and biofilm phases. Ecol Evol. 2014;4:4444–53.

    PubMed  PubMed Central  Article  Google Scholar 

  84. Pernthaler J. Predation on prokaryotes in the water column and its ecological implications. Nat Rev Microbiol. 2005;3:537–46.

    CAS  PubMed  Article  Google Scholar 

  85. Muller H. The relative importance of different ciliate taxa in the pelagic food web of lake constance. Microb Ecol. 1989;18:261–73.

    CAS  PubMed  Article  Google Scholar 

  86. Boenigk J, Arndt H. Bacterivory by heterotrophic flagellates: community structure and feeding strategies. Antonie Van Leeuwenhoek. 2002;81:465–80.

    PubMed  Article  Google Scholar 

  87. Battistini R, Marcucci E, Verani M, Di Giuseppe G, Dini F, Carducci A. Ciliate-adenovirus interactions in experimental co-cultures of Euplotes octocarinatus and in wastewater environment. Eur J Protistol. 2013;49:381–8.

    PubMed  Article  Google Scholar 

  88. Scheid P, Schwarzenberger R. Acanthamoeba spp. as vehicle and reservoir of adenoviruses. Parasitol Res. 2012;111:479–85.

    PubMed  Article  Google Scholar 

  89. Rigotto C, Hanley K, Rochelle PA, De Leon R, Barardi CR, Yates MV. Survival of adenovirus types 2 and 41 in surface and ground waters measured by a plaque assay. Environ Sci Technol. 2011;45:4145–50.

    CAS  PubMed  Article  Google Scholar 

  90. Ibrahim EME, El-Liethy MA, Abia ALK, Hemdan BA, Shaheen MN. Survival of E. coli O157:H7, Salmonella Typhimurium, HAdV2 and MNV-1 in river water under dark conditions and varying storage temperatures. Sci Total Environ. 2019;648:1297–304.

    CAS  PubMed  Article  Google Scholar 

  91. Rachmadi AT, Kitajima M, Pepper IL, Gerba CP. Enteric and indicator virus removal by surface flow wetlands. Sci Total Environ. 2016;542:976–82.

    CAS  PubMed  Article  Google Scholar 

  92. Pinheiro MD, Power ME, Butler BJ, Dayeh VR, Slawson R, Lee LE, Lynn DH, Bols NC. Use of Tetrahymena thermophila to study the role of protozoa in inactivation of viruses in water. Appl Environ Microbiol. 2007;73:643–9.

    CAS  PubMed  Article  Google Scholar 

  93. Deng L, Krauss S, Feichtmayer J, Hofmann R, Arndt H, Griebler C. Grazing of heterotrophic flagellates on viruses is driven by feeding behaviour. Environ Microbiol Rep. 2014;6:325–30.

    CAS  PubMed  Article  Google Scholar 

  94. Hennemuth W, Rhoads LS, Eichelberger H, Watanabe M, Van Bell KM, Ke L, Kim H, Nguyen G, Jonas JD, Veith D, Van Bell CT. Ingestion and inactivation of bacteriophages by Tetrahymena. J Eukaryot Microbiol. 2008;55:44–50.

    PubMed  Article  Google Scholar 

  95. Booncharoen N, Mongkolsuk S, Sirikanchana K. Comparative persistence of human sewage-specific enterococcal bacteriophages in freshwater and seawater. Appl Microbiol Biotechnol. 2018;102:6235–46.

    CAS  PubMed  Article  Google Scholar 

  96. Yang Y, Griffiths MW. Comparative persistence of subgroups of F-specific RNA phages in river water. Appl Environ Microbiol. 2013;79:4564–7.

    CAS  PubMed  PubMed Central  Article  Google Scholar 

  97. Pirlot S, Unrein F, Descy JP, Servais P. Fate of heterotrophic bacteria in Lake Tanganyika (East Africa). FEMS Microbiol Ecol. 2007;62:354–64.

    CAS  PubMed  Article  Google Scholar 

  98. Servais P, Garcia-Armisen T, George I, Billen G. Fecal bacteria in the rivers of the Seine drainage network (France): Sources, fate and modelling. Sci Total Environ. 2007;375:152–67.

    CAS  PubMed  Article  Google Scholar 

  99. Zingel P, Agasild H, Noges T, Kisand V. Ciliates are the dominant grazers on pico- and nanoplankton in a shallow, naturally highly eutrophic lake. Microb Ecol. 2007;53:134–42.

    PubMed  Article  Google Scholar 

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Acknowledgements

Authors would like to thank US Army Corps of Engineers, Louisville District, for access to William H. Harsha lake field site.

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The United States Environmental Protection Agency through its Office of Research and Development funded and managed the research described here. It has been subjected to Agency’s administrative review and approved for publication. The views expressed in this article are those of the author(s) and do not necessarily represent the views or policies of the U.S. Environmental Protection Agency. Mention of trade names or commercial products does not constitute endorsement or recommendation for use.

Funding

All funding was received through EPA Safe and Sustainable Water Resources (SSWR, task 3.02a).

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  1. United States Environmental Protections Agency, Cincinnati, OH, 45268, USA

    Brian R. McMinn, Eric R. Rhodes, Emma M. Huff & Asja Korajkic

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  1. Brian R. McMinn
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  2. Eric R. Rhodes
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Contributions

Study design (AK, BRM, ERR), field work (AK, BRM, ERR), laboratory work (AK, BRM, ERR, EMH), data analyses (AK), manuscript preparation (AK, BRM), final manuscript proof-reading and editing (AK, BRM, ERR, EMH). All authors read and approved the final manuscript.

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Correspondence to Asja Korajkic.

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McMinn, B.R., Rhodes, E.R., Huff, E.M. et al. Decay of infectious adenovirus and coliphages in freshwater habitats is differentially affected by ambient sunlight and the presence of indigenous protozoa communities. Virol J 17, 1 (2020). https://doi.org/10.1186/s12985-019-1274-x

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  • DOI: https://doi.org/10.1186/s12985-019-1274-x

Keywords

  • Coliphage
  • Adenovirus
  • Decay
  • Sunlight
  • Protozoa

Virology Journal

ISSN: 1743-422X