International Journal of Zoology
Volume 2020 (2020), Article ID 4731686, 7 pages
https://doi.org/10.1155/2020/4731686

Research Article

Use of Environmental DNA to Determine Fantail Darter (Etheostoma flabellare) Density in a Laboratory Setting: Effects of Biomass and Filtration Method

Ramon A. Guivas and Ben F. Brammell

Department of Science and Health, Asbury University, Wilmore, KY 40390, USA

Correspondence should be addressed to Ben F. Brammell; ben.brammell@asbury.edu

Received 9 December 2019; Accepted 23 March 2020; Published 17 June 2020

Academic Editor: Andrea Galimberti

Copyright © 2020 Ramon A. Guivas and Ben F. Brammell. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract.

Estimating fish abundance/biomass holds great importance for freshwater ecology and fisheries management, but current techniques can be expensive, time-consuming, and potentially harmful to target organisms. Environmental DNA (eDNA) has proven an effective and efficient technique for presence/absence detection of freshwater vertebrates. Additionally, recent studies report correlations between target organism density/biomass and eDNA levels, although widespread application of this technique is limited by the number of studies examining this relationship in various species and settings. Additionally, filter clogging is a commonly encountered issue in eDNA studies in environments with significant sediment and/or phytoplankton algae. Frequently, a sample must be split into multiple aliquots and filtered separately in order to process the entire sample. The present study examines both the relationship between biomass and eDNA and the effects of single versus multiple filter sampling on eDNA concentrations of fantail darters (Etheostoma flabellare) in a laboratory setting. Tank tests were performed in quadruplicate at four environmentally relevant fantail biomass levels. eDNA samples were collected and processed in parallel (one as a whole through a single filter and one in parts through multiple filters). Species-specific primers and a probe were developed for E. flabellare from cytochrome b sequences obtained from locally collected specimens, and real-time quantitative PCR was used to analyze eDNA levels at each biomass. Significant correlations were observed with increasing biomass for both methods, although this relationship was stronger for samples processed by the multiple filter method. These data should be useful in eDNA studies in which turbidity necessitates the use of multiple filters per sample as well as in the use of eDNA to estimate darter populations.

1. Introduction

Since its inception in macroinvertebrate studies in 2008 [1], environmental DNA (eDNA) has become firmly established as an effective method [2–4] and holds great promise for increasing the ease and scope of ecological studies. In aquatic systems, single species eDNA detection has primarily been applied to invasive species monitoring and conservation of threatened or endangered species [5]. Invasive species monitored using eDNA include Asian carp (Hypophthalmichthys sp.) [6] and multiple species of molluscs [7–9], crustaceans [10–12], amphibians [1, 13, 14], and reptiles [15, 16]. Rare and endangered taxa eDNA monitoring has included species such as Chinook salmon (Oncorhynchus tshawytscha) [17], eastern hellbenders (Cryptobranchus alleganiensis) [18, 19], great crested newts (Triturus cristatus) [20], the Japanese crayfish (Cambaroides japonicas) [21], and the Trinidad golden tree frog (Phytotriades auratus) [22], among others. Single species presence/absence eDNA monitoring is a well-established and widely used technique, in some cases enabling detection of cryptic species where traditional methods were unsuccessful [23, 24].

Recent studies also reveal correlations between organismal abundance and eDNA levels [25–28], indicating the possibility of quantitative eDNA analysis for population assessment. Laboratory studies with jack mackerels (Trachurus japonicus) [29], adult sea lampreys (Petromyzon marinus) [30], round gobies (Neogobius melanostomus) [31], and common carp (Cyprinus carpio) [25] all report positive correlations between eDNA levels and fish density or biomass. However, larval lamprey densities showed no correlation with eDNA in tank experiments [30]. Field studies examining the use of eDNA to quantify tadpole populations of the stream dwelling frog Odorrana splendida report a significant, but not strong, correlation between biomass and eDNA levels [27]. A study examining spawning salmon abundance and eDNA found eDNA levels varied with salmon density as well as numerous other factors [28]. Likewise, Lacoursière-Roussel et al. [32] reported a positive but weak correlation between catch per unit effort and lake trout (Salvelinus namaycush) eDNA levels. Additionally, numerous studies have observed seasonal fluctuations in eDNA, which are often believed to be associated with reproductive behavior [18, 28, 33]. While an emerging understanding supports the quantitative use of eDNA in population assessment, many factors appear to influence eDNA release [34] and persistence [35] in the environment, and additional studies are needed to validate eDNA as a density detection tool across various habitats and species [28, 36, 37].

Although quality assurance guidelines concerning eDNA collection and extraction have been established [38], numerous sample collection and preservation techniques exist, and the technique has been demonstrated to impact detection probability [39–43]. Although both precipitation and filtration have been used to capture DNA, filtration has been demonstrated to capture more eDNA from water samples [39, 41, 42] and as such is currently overwhelmingly the method of choice in eDNA studies. The clogging of filters by phytoplankton or suspended sediment is a frequently encountered issue in eDNA collection [25, 38, 44], particularly in turbid, lentic waters [45]. Filter clogging is typically countered by prefiltering [25], using larger filter pore sizes [25], or by dividing the sample and utilizing several filters to minimize clogging [19, 38]. Both prefiltering [46] and increasing filter pore size [42, 47] have been shown to reduce eDNA recovery. Although dividing a sample among several filters is a frequently employed technique [19], we are unaware of any studies examining its impact on eDNA efficacy or detection probability. The present study examines the hypothesis that multiple, as opposed to single filter, processing of water samples maintains or improves eDNA efficacy.

Darters are classified as Etheostomatinae, a subclade of Percidae that contains approximately 250 species endemic to eastern North America, comprising more than 20% of North America ichthyofauna [48]. Significant anatomical and physiological differences exist between darters and other members of Percidae [49, 50]. Darters are frequently in need of study as threatened or endangered species; currently, 27 darter species are listed as federally threatened or endangered on the U.S. Endangered Species List. Fantail darters (Etheostoma flabellare), a small, stream dwelling darter, common throughout much of their range in eastern North America [51, 52], were selected for this study as easily accessible members of this wide ranging subclade. We are aware of only one other darter eDNA study, in which eDNA was used to detect the presence of the federally endangered slackwater darter (Etheostoma boschungi) [53]. The present study provides data useful to approaching darter eDNA density studies.

eDNA offers great promise as a compliment [54] and perhaps in some cases a substitute for traditional field studies where such studies are difficult or impractical. This study provides novel data concerning two parameters of great significance to the application of eDNA in field ecology studies:

(1)The effect of increasing target organism density on eDNA levels(2)The effect of single versus multiple filters to quantitatively assess eDNA concentrations

2. Methods

2.1. Fish Collection and Housing

Fantail darters (Etheostoma flabellare) were collected from Jessamine Creek (Jessamine County, Kentucky) via electrofishing (KYDFW Permit#1811153) and acclimated to lab conditions in aged tap water over seven days prior to transfer to experimental tanks. Initial temperature of holding water in the lab matched stream temperature (6°C) at the time of collection. Water temperature was increased 2°C a day for seven days until it reached 20°C, and water was maintained at 20°C throughout the remainder of the holding period and the experiment. Fish were housed in a 208-liter tank prior to the experiment and fed commercially purchased blood worms ad libitum. All applicable international, national, and institutional guidelines for the ethical care and use of fish were followed.

2.2. eDNA Trials

Four environmentally relevant fantail darter biomass levels were calculated based on published studies of observed fantail darter densities [55, 56]. Tank tests for each of the four biomass levels were performed in quadruplicate; a negative control tank was included for each level to assess potential cross contamination between tanks. Experimental tanks (38 L) were held in an environmental chamber under regulated conditions (12 h day/12 h night, 20°C, aeration) for 96 hours, and fish were fed blood worms ad libitum during the course of the experiment.

2.3. eDNA Collection

Takahara et al. [25] and Maruyama et al. [57] both reported that eDNA initially spiked and then reached equilibrium near day four in fish tank tests, so a 96-hour end point was utilized for the present study. At 96 hours, all darters were removed, and each tank was homogenized by stirring for 1 minute to homogenously distribute DNA. Two one liter water samples were collected from each tank using 3.8-liter high-density polyethylene containers not previously exposed to animal DNA and thoroughly washed with bleach and rinsed with distilled water. Water samples were processed through a 47 mm diameter glass microfiber filter (VWR, 0.42 mm thickness and 0.7 μm pore size) in a manner similar to previous studies [6, 42, 58] with vacuum filtration immediately using one of the two methods: one sample from each tank was filtered through a single filter and the other was divided into three 333 ml aliquots, each of which was filtered separately. Filters were extracted immediately.

2.4. DNA Extraction

eDNA extraction was performed using a DNeasy blood and tissue kit (Qiagen), demonstrated to provide superior yields relative to other extraction methods [59], and a modified version of a published protocol [54]. Briefly, whole filters were cut into 30–40 pieces and incubated at 56°C overnight in 720 μl ATL buffer and 80 μl Proteinase K. Final elutions were performed twice into 400 μl of AE buffer, and the extracted DNA was stored at −20°C until analysis.

Tissue DNA extraction was performed using a DNeasy blood and tissue kit (Qiagen) according to the provided protocol. Tissue was lysed overnight at 56°C in proteinase K and eluted twice to increase DNA yield.

2.5. Primer Design

Cytochrome b was sequenced (GenBank: KT880219.1) from locally collected fantail darters (Jessamine Creek, Jessamine County, KY: 37.859380, −84.630755) using published primers [60]. Using this sequence, a species-specific primer probe assay was designed that amplifies a 118 bp amplicon within E. flabellare cytochrome b:

 Forward primer: 5′-AAGCGAAGAAGCGAGTTAGG-3′ Reverse primer: 5′-GGTGCTACGGTCATCACTAATC-3′ Probe: 5′-6 FAM/CCCACATAA/ZEN/G GCACTGCAGAGAGT/3IABkFQ-3′

Three other species of the family Percidae occurring sympatrically with E. flabellare in mid-sized central Kentucky streams were considered in the design of the primers and probes. We sequenced cytochrome b from locally collected (Jessamine Creek, Jessamine County, KY: 37.859380, −84.630755) sympatric specimens using the same primer set [60] and designed primers and probes with a minimum of three mismatches in each oligonucleotide (f. primer, r. primer, and probe) based on reported requirements for specificity [61]. Sympatric species considered include Etheostoma caeruleum (KT880220.1), Etheostoma blennioides (KT880218.1), and Percina caprodes (KT880217.1). Nonspecific amplification was tested using tissue extracted DNA from each of these species.

2.6. eDNA Analysis

Dilution of tank-extracted samples showed evidence of inhibition based on reduced product DNA yields despite the use of TaqMan Environmental Master Mix 2.0. Nondiluted samples amplified 9.8% of the DNA yield observed at the 1 : 20 dilution that was used for analysis (data available in Appendix A). A 1 : 20 dilution was selected based on previous studies indicating optimal yields observed at 1 : 20 dilutions [16] and the lower biomass to water ratios utilized in the present study relative to that study. Environmental DNA was quantified using a StepOnePlusTM Real-Time PCR system (Life Technologies, Carlsbad, CA, USA) in optical 96-well PCR plates. Each plate contained no fish control tank samples to assess contamination and fantail tissue samples as a positive control. Each 20 μl reaction contained the following: TaqMan EMM (10 μL), nuclease free water (7 μL), eDNA extract (2 μL), and E. flabellare primer/probe mix (1 μL). Thermocycler conditions were as follows: 95°C for 10 min, 55 cycles of 95°C for 15 s, and 60°C for one minute.

DNA extractions from E. flabellare fin clips were used to generate a standard as standards for the qPCR analysis. We used a Qubit 4 Fluorometer (Thermo Fisher) and a Qubit 1X dsDNA HS Assay Kit (Cat. No. Q32850) to quantify the DNA concentrations from the tissue extract. We then diluted a 7.88 ng/μl fantail darter DNA extract to five levels to use as a standard curve: 0.5 pg/l, 5 pg/l, 50 pg/l, 500 pg/l, and 5,000 pg/l. These dilutions cover the range of DNA concentrations that were observed in tank water-extracted eDNA in this study.

2.7. Statistical Analysis

Type I linear regression was used to examine the relationship of darter biomass and eDNA concentration within a single filter method following the previously published methods [44]. Both eDNA concentration and fish biomass were log transformed following the previously published methods [44, 62]. All tests were performed using IBM SPSS Statistics 25.

3. Results

Data from the single filter method trial indicated a positive correlation between darter biomass and eDNA concentration (Figure 1(a), , r² = 0.08). Interestingly, the multiple filter method demonstrated a significant () and stronger correlation (r² = 0.30) between these same two variables in parallel analysis (Figure 1(b)).

(a)

(b)

Figure 1: eDNA concentrations at various biomass levels for single filter method () (a) and multiple filter method () (b).

Amplification was observed in every tank containing darters, but no amplification was observed in any of the no fish control tanks, included in each trial to assess potential transfer of DNA between tanks. PCR efficiency was 97.2% as determined using a standard curve of diluted E. flabellare tissue.

4. Discussion

The few laboratory studies that have been completed examining eDNA and biomass utilize biomass ranges up to 200-fold, much greater than the more environmentally realistic 2-, 4-, and 8-fold differences examined in the present study. In tank studies (200 L) with Japanese mackerel (Trachurus japonicus) with an average of 6.9 g, 40.2 g, and 319.5 g of total biomass per tank, eDNA shedding rates differed between all groups at 13°C but not at 18, 23, or 28°C [29]. The treatments in this study represent a 6-fold and 46-fold increase in biomass in the upper two treatments compared to the lower. Trials with sea lampreys (Petromyzon marinus) reported significant increases in biomass with 0, 2, 20, and 200 adults per 2000 L tank but not with 0, 1, 5, and 25 larva in 28  L aquaria [30]. Assuming homogenous weight of the adult sea lampreys, these treatments exceeded biomass levels in the present study in all but the lowest treatment. Juvenile carp (Cyprinus carpio) trials in 9 L tanks found significant correlations between 16 g, 79 g (5-fold), and 158 g (10-fold) biomass and eDNA levels [25], representing similar biomass level treatments to those utilized in the present experiment. The results of the present experiment represent the lowest biomass/eDNA relationships tested to date and are consistent with the results of previous studies, most of which reported positive correlations between biomass and eDNA.

The relationship between biomass and eDNA levels was significant and positive for both filter methods, although the r² values were not large (0.29 and 0.49). These values are similar to those observed by Iwai et al. [27]. When comparing tadpole abundance (0.32) and biomass (0.29) to eDNA copy number in lotic samples sites, they observed large variations in the biomass-eDNA relationship within a single stream. Likewise, Baldigo et al. [44] reported r² values of 0.44 (density) and 0.25 (biomass) in Adirondack mountain headwater brook trout (Salvelinus fontinalis) populations. These data support a growing data set indicating a loose correlation between organism abundance and eDNA concentrations.

The results of the present study support the premise that multiple filters improve the resolving capacity of eDNA as the combined filter method (333 ml per filter) demonstrated a significant relationship between biomass and eDNA, while the same samples processed in a single filter (1 L per filter) did not. Hunter et al. [63] report a 10% increase in DNA yield utilizing multiple as opposed to single filters; although this was not visible in our study, it is possible that a larger sample size would have revealed a similar trend. Filtering larger volumes of water leads to a greater concentration of inhibitors [64], which perhaps could be linked to greater inconsistency in DNA detection in single filter samples if inhibitors are removed more effectively in the multiple filter process as less inhibitor would be present in each filter during the extraction process. Regardless of the mechanism, these results provide evidence that the use of multiple filters to avoid the frequently encountered problem of filter clogging [65–67] is acceptable and perhaps beneficial.

The positive correlation between biomass and eDNA has been previously observed, but not in a small, stream dwelling species at environmentally relevant densities. Doi et al. [26] reported positive correlations between Plecoglossus altivelis densities as estimated from snorkeling surveys and eDNA levels. Klymus et al. [68] observed a positive correlation in eDNA and biomass in two species of bighead carp (Hypophthalmichthys nobilis and Hypophthalmichthys molitrix). Our work builds on these previous efforts by comparing environmentally relevant biomass levels with eDNA concentrations and providing data supporting the validity of utilizing multiple, versus single, filters in field studies.

Data Availability

The data used to support the findings of this study are available upon request to the corresponding author.

Conflicts of Interest

The authors declare that they have no conflicts of interest.

Acknowledgments

This project was supported by an Undergraduate Research Grant (#9118243-Guivas) from the Kentucky Academy of Science.

Supplementary Materials

Table 1: nondiluted eDNA concentrations for nondiluted samples. These data were not used in the publication because they appear to be inhibited relative to 1 : 20 diluted samples. Table 2: eDNA concentrations for samples diluted 1 : 20 following DNA extraction. Table 3: CT values for samples following a 1 : 20 dilution of extracted DNA. Table 4: CT values for nondiluted samples. Figure 1: eDNA concentrations at various biomass levels for single filter method (A) () and multiple filter method (B) () for nondiluted samples. These data were not used in the paper because they appear to be inhibited relative to 1 : 20 diluted samples. (Supplementary Materials)

References

G. F. Ficetola, C. Miaud, F. Pompanon, and P. Taberlet, “Species detection using environmental DNA from water samples,” Biology Letters, vol. 4, no. 4, pp. 423–425, 2008.

N. A. Sawaya, A. Djurhuus, C. J. Closek et al., “Assessing eukaryotic biodiversity in the Florida keys national marine sanctuary through environmental DNA metabarcoding,” Ecology and Evolution, vol. 9, no. 3, pp. 1029–1040, 2019.

Q. Mauvisseau, A. Coignet, C. Delaunay, F. Pinet, D. Bouchon, and C. Souty-Grosset, “Environmental DNA as an efficient tool for detecting invasive crayfishes in freshwater ponds,” Hydrobiologia, vol. 805, no. 1, pp. 163–175, 2018.

J. C. Dysthe, T. W. Franklin, K. S. McKelvey, M. K. Young, and M. K. Schwartz, “An improved environmental DNA assay for bull trout (Salvelinus confluentus) based on the ribosomal internal transcribed spacer I,” PLoS One, vol. 13, no. 11, Article ID e0206851, 2018.

C. I. M. Adams, L. A. Hoekstra, M. R. Muell, and F. J. Janzen, “A brief review of non-avian reptile environmental DNA (eDNA), with a case study of painted turtle (Chrysemys picta) eDNA under field conditions,” Preprints, vol. 11, p. 50, 2019.

C. L. Jerde, A. R. Mahon, W. L. Chadderton, and D. M. Lodge, ““Sight-unseen” detection of rare aquatic species using environmental DNA,” Conservation Letters, vol. 4, no. 2, pp. 150–157, 2011.

C. S. Goldberg, A. Sepulveda, A. Ray, J. Baumgardt, and L. P. Waits, “Environmental DNA as a new method for early detection of New Zealand mudsnails (Potamopyrgus antipodarum),” Freshwater Science, vol. 32, no. 3, pp. 792–800, 2013.

K. E. Klymus, N. T. Marshall, and C. A. Stepien, “Environmental DNA (eDNA) metabarcoding assays to detect invasive invertebrate species in the Great Lakes,” PLoS One, vol. 12, no. 5, Article ID e0177643, 2017.

Z. Xia, A. Zhan, Y. Gao, L. Zhang, G. D. Haffner, and H. J. MacIsaac, “Early detection of a highly invasive bivalve based on environmental DNA (eDNA),” Biological Invasions, vol. 20, no. 2, pp. 437–447, 2018.

K. J. Carim, K. R. Christianson, K. M. McKelvey et al., “Environmental DNA marker development with sparse biological information: a case study on opossum shrimp (Mysis diluviana),” PLoS One, vol. 11, no. 8, Article ID e0161664, 2016.

M. M. Dougherty, E. R. Larson, M. A. Renshaw et al., “Environmental DNA (eDNA) detects the invasive rusty crayfish Orconectes rusticus at low abundances,” Journal of Applied Ecology, vol. 53, no. 3, pp. 722–732, 2016.

T. Forsström and A. Vasemägi, “Can environmental DNA (eDNA) be used for detection and monitoring of introduced crab species in the Baltic Sea?” Marine Pollution Bulletin, vol. 109, no. 1, pp. 350–355, 2016.

T. Dejean, A. Valentini, C. Miquel, P. Taberlet, E. Bellemain, and C. Miaud, “Improved detection of an alien invasive species through environmental DNA barcoding: the example of the American bullfrog Lithobates catesbeianus,” Journal of Applied Ecology, vol. 49, no. 4, pp. 953–959, 2012.

J. Secondi, T. Dejean, A. Valentini, B. Audebaud, and C. Miaud, “Detection of a global aquatic invasive amphibian, Xenopus laevis, using environmental DNA,” Amphibia-Reptilia, vol. 37, no. 1, pp. 131–136, 2016.

A. J. Piaggio, R. M. Engeman, M. W. Hopken et al., “Detecting an elusive invasive species: a diagnostic PCR to detect Burmese python in Florida waters and an assessment of persistence of environmental DNA,” Molecular Ecology Resources, vol. 14, no. 2, pp. 374–380, 2014.

C. M. Davy, A. G. Kidd, and C. C. Wilson, “Development and validation of environmental DNA (eDNA) markers for detection of freshwater turtles,” PLoS One, vol. 10, no. 7, Article ID e0130965, 2015.

M. B. Laramie, D. S. Pilliod, and C. S. Goldberg, “Characterizing the distribution of an endangered salmonid using environmental DNA analysis,” Biological Conservation, vol. 183, pp. 29–37, 2015.

S. F. Spear, J. D. Groves, L. A. Williams, and L. P. Waits, “Using environmental DNA methods to improve detectability in a hellbender (Cryptobranchus alleganiensis) monitoring program,” Biological Conservation, vol. 183, pp. 38–45, 2015.

A. J. Santas, T. Persaud, B. A. Wolfe, and J. M. Bauman, “Noninvasive method for a statewide survey of eastern hellbenders Cryptobranchus alleganiensis using environmental DNA,” International Journal of Zoology, vol. 2013, Article ID 174056, 6 pages, 2013.

H. C. Rees, C. A. Baker, D. S. Gardner, B. C. Maddison, and K. C. Gough, “The detection of great crested newts year round via environmental DNA analysis,” BMC Research Notes, vol. 10, no. 1, 2017.

K. Ikeda, H. Doi, K. Tanaka, T. Kawai, and J. N. Negishi, “Using environmental DNA to detect an endangered crayfish Cambaroides japonicus in streams,” Conservation Genetics Resources, vol. 8, no. 3, pp. 231–234, 2016.

S. Brozio, C. Manson, E. Gourevitch et al., “Development and application of an eDNA method to detect the critically endangered Trinidad golden tree frog (Phytotriades auratus) in bromeliad phytotelmata,” PLoS One, vol. 12, no. 2, Article ID e0170619, 2017.

L. M. Gargan, T. Morato, C. K. Pham, J. A. Finarelli, J. E. L. Carlsson, and J. Carlsson, “Development of a sensitive detection method to survey pelagic biodiversity using eDNA and quantitative PCR: a case study of devil ray at seamounts,” Marine Biology, vol. 164, no. 5, 2017.

E. E. Sigsgaard, H. Carl, P. R. Møller, and P. F. Thomsen, “Monitoring the near-extinct European weather loach in Denmark based on environmental DNA from water samples,” Biological Conservation, vol. 183, pp. 46–52, 2015.

T. Takahara, T. Minamoto, H. Yamanaka, H. Doi, and Z. i. Kawabata, “Estimation of fish biomass using environmental DNA,” PLoS One, vol. 7, no. 4, Article ID e35868, 2012.

H. Doi, R. Inui, Y. Akamatsu et al., “Environmental DNA analysis for estimating the abundance and biomass of stream fish,” Freshwater Biology, vol. 62, no. 1, pp. 30–39, 2017.

N. Iwai, K. Yasumiba, and T. Takahara, “Efficacy of environmental DNA to detect and quantify stream tadpoles of Odorrana splendida,” Royal Society Open Science, vol. 6, no. 1, Article ID 181798, 2019.

M. D. Tillotson, R. P. Kelly, J. J. Duda, M. Hoy, J. Kralj, and T. P. Quinn, “Concentrations of environmental DNA (eDNA) reflect spawning salmon abundance at fine spatial and temporal scales,” Biological Conservation, vol. 220, pp. 1–11, 2018.

S. Yamamoto, T. Jo, H. Murakami, T. Minamoto, and R. Masuda, “Effect of water temperature and fish biomass on environmental DNA shedding, degradation, and size distribution,” Ecology and Evolution, vol. 9, no. 3, pp. 1135–1146, 2019.

N. Schloesser, “Correlating sea lamprey density with environmental DNA detections in the lab,” Management of Biological Invasions, vol. 9, no. 4, pp. 483–495, 2018.

M. B. Nevers, M. N. Byappanahalli, C. C. Morris et al., “Environmental DNA (eDNA): a tool for quantifying the abundant but elusive round goby (Neogobius melanostomus),” PLoS One, vol. 13, no. 1, Article ID e0191720, 2018.

A. Lacoursière-Roussel, G. Côté, V. Leclerc, and L. Bernatchez, “Quantifying relative fish abundance with eDNA: a promising tool for fisheries management,” Journal of Applied Ecology, vol. 53, no. 4, pp. 1148–1157, 2016.

L. S. De Souza, J. C. Godwin, M. A. Renshaw, and E. Larson, “Environmental DNA (eDNA) detection probability is influenced by seasonal activity of organisms,” PLoS One, vol. 11, no. 10, Article ID e0165273, 2016.

R. Lance, K. Klymus, C. Richter et al., “Experimental observations on the decay of environmental DNA from bighead and silver carps,” Management of Biological Invasions, vol. 8, no. 3, pp. 343–359, 2017.

M. A. Barnes, C. R. Turner, C. L. Jerde, M. A. Renshaw, W. L. Chadderton, and D. M. Lodge, “Environmental conditions influence eDNA persistence in aquatic systems,” Environmental Science & Technology, vol. 48, no. 3, pp. 1819–1827, 2014.

A. S. Buxton, J. J. Groombridge, and R. A. Griffiths, “Is the detection of aquatic environmental DNA influenced by substrate type?” PLoS One, vol. 12, no. 8, Article ID e0183371, 2017.

M. A. Barnes and C. R. Turner, “The ecology of environmental DNA and implications for conservation genetics,” Conservation Genetics, vol. 17, no. 1, pp. 1–17, 2016.

C. S. Goldberg, C. R. Turner, K. Deiner et al., “Critical considerations for the application of environmental DNA methods to detect aquatic species,” Methods in Ecology and Evolution, vol. 7, no. 11, pp. 1299–1307, 2016.

M. P. Piggott, “Evaluating the effects of laboratory protocols on eDNA detection probability for an endangered freshwater fish,” Ecology and Evolution, vol. 6, no. 9, pp. 2739–2750, 2016.

M. Majaneva, O. H. Diserud, S. H. C. Eagle, E. Boström, M. Hajibabaei, and T. Ekrem, “Environmental DNA filtration techniques affect recovered biodiversity,” Scientific Reports, vol. 8, no. 1, 2018.

K. Deiner, J.-C. Walser, E. Mächler, and F. Altermatt, “Choice of capture and extraction methods affect detection of freshwater biodiversity from environmental DNA,” Biological Conservation, vol. 183, pp. 53–63, 2015.

J. J. Eichmiller, L. M. Miller, and P. W. Sorensen, “Optimizing techniques to capture and extract environmental DNA for detection and quantification of fish,” Molecular Ecology Resources, vol. 16, no. 1, pp. 56–68, 2016.

M. A. Renshaw, B. P. Olds, C. L. Jerde, M. M. Mcveigh, and D. M. Lodge, “The room temperature preservation of filtered environmental DNA samples and assimilation into a phenol-chloroform-isoamyl alcohol DNA extraction,” Molecular Ecology Resources, vol. 15, no. 1, pp. 168–176, 2015.

B. P. Baldigo, L. A. Sporn, S. D. George, and J. A. Ball, “Efficacy of environmental DNA to detect and quantify brook trout populations in headwater streams of the Adirondack mountains, New York,” Transactions of the American Fisheries Society, vol. 146, no. 1, pp. 99–111, 2017.

L. R. Harper, A. S. Buxton, H. C. Rees et al., “Prospects and challenges of environmental DNA (eDNA) monitoring in freshwater ponds,” Hydrobiologia, vol. 826, no. 1, pp. 25–41, 2019.

C. R. Turner, M. A. Barnes, C. C. Y. Xu, S. E. Jones, C. L. Jerde, and D. M. Lodge, “Particle size distribution and optimal capture of aqueous macrobial eDNA,” Methods in Ecology and Evolution, vol. 5, no. 7, pp. 676–684, 2014.

Z. Liang and A. Keeley, “Filtration recovery of extracellular DNA from environmental water samples,” Environmental Science & Technology, vol. 47, no. 16, pp. 9324–9331, 2013.

T. J. Near, C. M. Bossu, G. S. Bradburd et al., “Phylogeny and temporal diversification of darters (Percidae: Etheostomatinae),” Systematic Biology, vol. 60, no. 5, pp. 565–595, 2011.

R. C. Cashner, D. E. Etnier, and W. C. Starnes, “The fishes of Tennessee,” Copeia, vol. 1995, no. 2, p. 508, 1995.

L. M. Page and L. E. Cordes, “Variation and systematics of etheostoma euzonum, the Arkansas saddled darter (pisces: Percidae),” Copeia, vol. 1983, no. 4, p. 1042, 1983.

R. A. Kuehne and R. W. Barbour, “The American darters,” Copeia, vol. 1984, no. 2, p. 562, 1984.

R. A. Knapp and R. C. Sargent, “Egg-mimicry as a mating strategy in the fantail darter, Etheostoma flabellare: females prefer males with eggs,” Behavioral Ecology and Sociobiology, vol. 25, no. 5, pp. 321–326, 1989.

A. M. Janosik and C. E. Johnston, “Environmental DNA as an effective tool for detection of imperiled fishes,” Environmental Biology of Fishes, vol. 98, no. 8, pp. 1889–1893, 2015.

C. S. Goldberg, D. S. Pilliod, R. S. Arkle, and L. P. Waits, “Molecular detection of vertebrates in stream water: a demonstration using rocky mountain tailed frogs and Idaho giant salamanders,” PLoS One, vol. 6, no. 7, Article ID e22746, 2011.

N. D. D. Mundahl and C. G. G. Ingersoll, “Early autumn movements and densities of Johnny (Etheostoma nigrum) and fantail (E. flabellare) darters in a southwestern Ohio stream,” The Ohio Journal of Science, vol. 43, no. 3, pp. 103–108, 1983.

K. Lindström and R. C. Sargent, “Food access, brood size and filial cannibalism in the fantail darter, Etheostoma flabellare,” Behavioral Ecology and Sociobiology, vol. 40, no. 2, pp. 107–110, 1997.

A. Maruyama, K. Nakamura, H. Yamanaka, M. Kondoh, and T. Minamoto, “The release rate of environmental DNA from juvenile and adult fish,” PLoS One, vol. 9, no. 12, Article ID e114639, 2014.

J. J. Eichmiller, P. G. Bajer, and P. W. Sorensen, “The relationship between the distribution of common carp and their environmental DNA in a small lake,” PLoS One, vol. 9, no. 11, Article ID e112611, 2014.

R. Hinlo, D. Gleeson, M. Lintermans, and E. Furlan, “Methods to maximise recovery of environmental DNA from water samples,” PLoS One, vol. 12, no. 6, Article ID e0179251, 2017.

T. R. Schmidt and J. R. Gold, “Complete sequence of the mitochondrial cytochrome b gene in the cherryfin shiner, Lythrurus roseipinnis (teleostei: cyprinidae),” Copeia, vol. 1993, no. 3, pp. 880–883, 1993.

T. M. Wilcox, K. S. McKelvey, M. K. Young et al., “Robust detection of rare species using environmental DNA: the importance of primer specificity,” PLoS One, vol. 8, no. 3, Article ID e59520, 2013.

P. F. Thomsen, J. Kielgast, L. L. Iversen et al., “Monitoring endangered freshwater biodiversity using environmental DNA,” Molecular Ecology, vol. 21, no. 11, pp. 2565–2573, 2012.

M. E. Hunter, J. A. Ferrante, G. Meigs-Friend, and A. Ulmer, “Improving eDNA yield and inhibitor reduction through increased water volumes and multi-filter isolation techniques,” Scientific Reports, vol. 9, no. 1, pp. 1–9, 2019.

J. Herder, A. Valentini, E. Bellemain et al., “Environmental DNA, A review of the possible applications for the detection of (invasive) species,” Tech. Rep., RAVON, Nijimegen, Netherlands, 2014, Technical report no. 2013-104.

A. Bruno, A. Sandionigi, A. Galimberti et al., “One step forwards for the routine use of high-throughput DNA sequencing in environmental monitoring. An efficient and standardizable method to maximize the detection of environmental bacteria,” Microbiologyopen, vol. 6, no. 1, Article ID e00421, 2017.

C. R. Turner, D. J. Miller, K. J. Coyne, and J. Corush, “Improved methods for capture, extraction, and quantitative assay of environmental DNA from Asian bigheaded carp (Hypophthalmichthys spp.),” PLoS One, vol. 9, no. 12, Article ID e114329, 2014.

K. J. Dunker, A. J. Sepulveda, R. L. Massengill et al., “Potential of environmental DNA to evaluate northern pike (Esox lucius) eradication efforts: an experimental test and case study,” PLoS One, vol. 11, no. 9, Article ID e0162277, 2016.

K. E. Klymus, C. A. Richter, D. C. Chapman, and C. Paukert, “Quantification of eDNA shedding rates from invasive bighead carp Hypophthalmichthys nobilis and silver carp Hypophthalmichthys molitrix,” Biological Conservation, vol. 183, pp. 77–84, 2015.