Etheostoma trisella
environmental DNA
loop-mediated isothermal amplification
freshwater
1 INTRODUCTION
The conservation of imperiled species requires an understanding of
current population stability and distribution. However, the surveillance
of aquatic species via the employment of traditional methods such as
netting and electrofishing are labor intensive, expensive, and
disruptive to the species of conservation concern (Jerde et al., 2011).
Many species that require monitoring are rare, and, in fact, the
probability of detecting aquatic taxa using traditional methods is low
when species density falls below a specific threshold (Dejean et al.,
2012). Furthermore, due to permitting restrictions, it is difficult to
rapidly assess populations of species protected by law. Some studies
have found that sampling methods such as electrofishing do not
accurately determine the abundance of targeted taxa despite their
invasive nature (Wilcox et al., 2016). Several eDNA studies report that
published methods did not capture the true diversity or abundance of
taxa known from traditional sampling (Cilleros et al., 2019; Drinkwater
et al., 2019). As such, a noninvasive, supplemental method is required
in order to monitor and provide protection to as many imperiled species
as possible.
Environmental DNA (eDNA) is the genetic material found in environmental
samples such as water, sediment, and air. Representing a conglomeration
of genetic material sourced to different organisms, eDNA molecules
originate from extracellular DNA (i.e., natural cell death followed by
cell lysis), cellular DNA (e.g., epithelial cells, mucus, and feces) or
microbial organisms (Díaz-Ferguson & Moyer, 2014; Nagler et al., 2018;
Pietramellara et al., 2009; Taberlet et al., 2012). DNA from
environmental sources was first utilized in the 1980’s for the
extraction of microbial DNA from sediment and later for bacterial
biomass assessment (Ogram et al., 1987; Paul et al., 1996).
Environmental DNA is now used in aquatic systems to detect invasive
species, monitor populations of threatened taxa, and trace fecal
contaminants (Díaz-Ferguson & Moyer, 2014; Goldberg et al., 2011; Jerde
et al., 2011; Layton et al., 2006).
The detection of target eDNA using traditional polymerase chain reaction
(PCR) and quantitative PCR (qPCR) at natural sites has shown limited
applicability in both freshwater and marine habitats (Foote et al.,
2012; Grey et al., 2018). In an effort to increase the utility of eDNA,
we have developed an eDNA detection method that applies loop-mediated
isothermal amplification (LAMP). Previous authors have prospected the
utility of LAMP for animal eDNA detection, but this study is the first
to demonstrate feasibility for freshwater fishes (Lee, 2017; Rees et
al., 2014). LAMP amplifies target sequences with high specificity by
initially binding four different primers to six complementary regions of
the target DNA; annealing of two primers to four target regions
maintains specificity as amplification progresses (Notomi et al., 2000).
This method uses a Bst DNA polymerase under isothermal conditions
to produce multiple stem-loop DNA structures (Notomi et al., 2000).
Furthermore, LAMP has the capability of amplifying small quantities of
target DNA with detection possible with as few as six copies in solution
during experimentation (Notomi et al., 2000).
While the use of genetic material isolated from nonliving sources for
the detection of targeted taxa is not novel, our custom molecular
methods are. We present here the application of our new method to the
surveillance of Etheostoma trisella (Percidae) populations.E. trisella , the Trispot Darter, is a small, freshwater fish
endemic to the Coosa River system of northwestern Georgia, southeastern
Tennessee, and northeastern Alabama (Page & Burr, 2011). The currently
understood range in Georgia and Tennessee includes the Conasauga River
system upstream of the confluence with the Coosawattee River including
Mill Creek and below Carter’s Lake, and the Oostanaula River between
Arumuchee Creek and Calhoun (Boschung & Mayden, 2004; Rare
Species Status Maps , 2020). In Alabama, E. trisella has recently
been recovered in Little Canoe Creek and Ballplay Creek
(FishMap.Org , 2018; O’Neil et al., 2009). Etheostoma
trisella occupies a specialized niche and migrates between two
seasonally interconnected habitats (Ryon, 1986). During the non-breeding
season (mid-April to mid-October), E. trisella populates
peripheral zones characterized by the presence of gravel, vegetation,
and a slow moving current. Seasonal rainfall allows the darter to
migrate upstream to the breeding grounds where they spawn from January
to March. This habitat consists of streams in seepage waters of pastures
and floodplains containing high amounts of vegetation. Given thatE. trisella has a severely limited distribution and is highly
impacted by anthropogenic activity, the darter was listed as
“Threatened” under the Endangered Species Act (ESA) in 2019
(Endangered and Threatened Wildlife and Plants; Threatened Species
Status for Trispot Darter , 2018). Therefore, surveillance of E.
trisella populations is essential in understanding how to best
implement conservation efforts.
2 MATERIALS AND METHODS
2.1 Sample Collection
We collected environmental water samples (n = 256) during 2019
(January-March) at 136 sites encompassing 17 watersheds in Georgia and
Alabama, USA (Figure 1). Collection sites were chosen to collectively
satisfy the following criteria: 1) localities of documented E.
trisella captures, 2) localities proximal to documented E.
trisella captures, and 3) localities where E. trisella have not
been documented. For each collection, two sealed 0.5L plastic water
bottles were emptied of the manufacturer’s contents and filled with
environmental water. Negative field controls were implemented as
suggested by Goldberg et al. (2016).
2.2 eDNA Extraction
We captured eDNA from environmental water samples by filtering through
Whatman Microfiber Filters (GE Healthcare, Chicago, IL, USA) housed in
Nalgene Analytical Test Filter Funnels (Thermo Fisher Scientific,
Waltham, MA, USA) using the Geopump Peristaltic Pump (Geotech
Environmental Equipment, Denver, CO, USA). Duplicate water samples were
combined during filtration. Filters containing residual particles were
cut into halves; one half was stored in 70% ethanol at 4 °C and the
other half used immediately in DNA extraction. DNA extraction was
performed using the Qiagen DNeasy Blood and Tissue Kit (Qiagen, Hilden,
Germany) and the QIAshredder (Qiagen, Hilden, Germany). DNA quality and
quantity were determined using standard gel electrophoresis on a 1.5%
agarose gel and a NanoDrop 2000 Spectrophotometer (Thermo Fisher
Scientific, Waltham, MA, USA). Purified eDNA was stored at 4 °C.
Filtration of water and extraction of eDNA were performed in a lab
environment cleaned with a 10% bleach solution.
2.3 eDNA Detection
We detected E. trisella presence by implementing a custom LAMP
protocol. LAMP primers were designed in PrimerExplorer (v. 5) within the
mitochondrial genome with specificity for the E. trisellacytochrome b (cyt b ) gene (Table 1) and optimized in a
temperature titration (LAMP Primer Designing
Software:PrimerExplorer , 2015; Sandel et al., 2020). Primer specificity
was verified using Primer-BLAST; our external primers alone yieldedE. trisella as the only expected target (Ye et al., 2012). LAMP
was performed with the following specifications suggested by the
manufacturer: 12.5µL LavaLAMP DNA Master Mix (2X; Lucigen, Middleton,
WI, USA), 2.5µL target-specific primer assay (Table 1), 1.0µL green
fluorescent dye (Lucigen, Middleton, WI, USA), bovine serum albumin (0.4
mg/mL; Thermo Fisher Scientific, Waltham, MA, USA), 7.5µL Invitrogen
UltraPure DNase/RNase-Free Distilled Water (Thermo Fisher Scientific,
Waltham, MA, USA), and 1.0µL template DNA for a total reaction volume of
25.0µL. Reactions were performed on a Stratagene Mx3000P (Agilent
Technologies, Santa Clara, CA, USA) under the following conditions: 4
min 90°C preheat, 30 min 74°C temperature hold, standard dissociation
curve, and 5 min 95°C enzyme deactivation. All reactions were performed
in triplicate including positive and negative controls. To ensure that
our primers amplify different genetic populations of E. trisella ,
we performed our LAMP protocol on a subset of fish representing three
genetically distinct populations (unpublished data). Although our
primers were designed to target E. trisella only, we determined
that two species of darter could potentially be cross-amplified. To
account for this possibility, we screened DNA samples from E.
jordani and E. rupestre , genetically similar species of darter
that may be detectable in our water samples.
Results were visualized using standard gel electrophoresis on a 1.5%
agarose gel, amplification plots and dissociation curves on the
Stratagene Mx3000P, and quantitative electrophoresis on a QIAxcel
Advanced System (Qiagen, Hilden, Germany). We performed quantitative
electrophoresis using the QIAxcel DNA High Resolution Kit (Qiagen,
Hilden, Germany) combined with a 15bp/600bp QX Alignment Marker (Qiagen,
Hilden, Germany) and 25-500bp QX DNA Size Marker (5ng/µL) under the
manufacturer’s suggested 0H800 method. A positive status was assigned to
experimental samples if they produced DNA segments comparable in size to
at least three of the five segments characterizing the positive control.
In addition, samples were only deemed positive upon reaching a positive
result in a second replicate (Mahon et al., 2013). A subset of positive
products was purified using the QIAquick Gel Extraction Kit (Qiagen,
Hilden, Germany) and sequenced with a SimpleSeq Kit (Eurofins Genomics,
Louisville, KY, USA) to verify template amplification. We used NCBI
BLAST to determine the origin of each DNA fragment (Altschul et al.,
1990). Statistical analyses were performed using the R Programming
Language including Fisher’s Exact Tests and covariance to determine
which factors may influence detection, if any (R Core Team, 2018).
3 RESULTS
After screening environmental water samples, 187 samples were found to
be negative for E. trisella presence and 69 positive (Figure 1).
At sites in Alabama, 24.5% of water samples screened positive (n = 151)
while 30.5% of samples in Georgia were positive (n = 105). Water
samples from different states were obtained by different groups of
collectors, and we show that detection of E. trisella was not
influenced by collector (p = 0.3911). However, the date of water
collection did influence detection (p < 0.05). Post hocpairwise analyses indicate that detections were lowest during the
beginning of our collection period (January) and increased in February
through March (p < 0.05). The sampling scheme (i.e.,
north/south or east/west) did not confound this temporal trend (Cov =
-0.11921).
Our primers amplified all genetic lineages of E. trisella tested.
With several mismatched sites in our primers, amplification of DNA
belonging to other Etheostoma species was not a source of false
positives as neither E. jordani nor E. rupestre amplified
using our E. trisella primer set. Sequence data confirmed
amplification of E. trisella DNA in a positive control and two
environmental water samples (Table 2). The positive control DNA sample
collected from E. trisella tissue matched across its entirety to
an E. trisella voucher specimen. The two eDNA fragments matchedE. trisella DNA across > 95.0% of nucleotides with
over 98.0% sequence coverage. One environmental sample however, did not
match to E. trisella and the source of the eDNA molecule(s)
sequenced could not be resolved. The sequence showed 91.49% nucleotide
identity to the bacterium Rahnella aquatilis , but at only 16.0%
coverage. In thirteen instances, environmental negative controls
screened positive. Attempts to extract positive bands from an agarose
gel for sequencing were inconclusive and did not provide any usable
data. The number of environmental negative controls that screened
positive did not show an association with collector (p = 0.3834).
4 DISCUSSION
Using noninvasive methods for the surveillance of imperiled species is
an attractive alternative to traditional methods, and we have presented
our contribution to this innovation. We screened 256 water samples from
Alabama and Georgia for E. trisella eDNA and were able to detect
their presence using our LAMP protocol. A comparable study was completed
that assessed E. trisella presence using eDNA and PCR in some of
the same Alabama streams which showed fewer positive detections (1.3%)
than we reported here (24.5%) (Johnston & Janosik, 2019).
In verifying that our field and laboratory protocols were sufficient to
detect E. trisella eDNA, we tested several positive controls
known to contain E. trisella DNA. Because our primer design was
broad enough to amplify DNA in positive control fish from different
genetic populations, we infer that our methods should be capable of
detecting fish from any of our localities. Including the proportion of
negative environmental controls, we were also able to rule out bias in
detection from the investigator (p = 0.3834). From these data we
conclude that all samples were handled in a similar fashion through the
point of collection, transport, and processing.
The protocol that we designed includes several checks to determine if
false positives are present. Our pipeline required negative and positive
controls as well as cleaning lab equipment with bleach solution as
described above. We also sequenced positive samples to verify the DNA
source. We detected E. trisella presence in 34.2% of the
environmental negative controls implemented (n = 38). We were unable to
isolate and sequence DNA fragments from these samples, but hypothesize
that amplification in negative controls may result from contamination or
amplification of eDNA from other sources. The incidence of false
positives in LAMP studies has been previously documented in disease
prevalence studies including that of malaria and thalassemia (Kollenda
et al., 2018; Wang et al., 2020). The addition of quenched fluorescent
primers to LAMP has been suggested to reduce the number of false
positives(Hardinge & Murray, 2019). Our reusable, bleached filter
funnels are a potential source of contamination, and closed filters are
an alternative that may reduce risks (Spens et al., 2017). Although our
primers were designed with specificity for E. trisella , the LAMP
protocol may cross-amplify DNA of other unintended targets. We
determined that our primers matched E. trisella along the
entirety of the cyt b gene and mismatched E. jordani andE. rupestre at multiple nucleotides (18 and 26 sites
respectively). We have shown that our primers show high specificity for
the intended target, and are not a likely source of false positives. We
postulate that environmental negative control water samples taken at
Shoal Creek on the Cahaba River screened positive due to contamination
or cross-amplification of a target not yet documented in genetic
databases. Inspection of the eDNA sequence that matched poorly toRahnella aquatilis showed no sites that were likely for binding
of our primers. We consider this sequence unresolved and not explicitly
identified as Rahnella aquatilis .
We stress that caution should be used when interpreting negative
results. Thus, the lack of detection at sites does not necessarily
indicate that E. trisella are not present at those localities,
but that our methods did not detect their presence. When utilizing eDNA,
false negatives may arise because template DNA is not amplified using
molecular methods despite presence of the target organism (Darling &
Mahon, 2011). Failure to amplify template DNA may occur because of DNA
degradation, low quality or quantity DNA, low primer specificity, PCR
inhibitors, and failed reactions (Klymus et al., 2015; Nathan et al.,
2015). In addition, replicates are required to accurately determine
diversity and detect rare species (Fonseca, 2018).
We report that detection probabilities varied significantly with the
date of collection, with the number of detections increasing at the end
of our collection period (i.e., expected end of spawning period). The
life history of E. trisella may play a particularly strong role
in why detections varied over time. E. trisella move between main
and spawning channels. Although we did sample from potential spawning
grounds throughout the expected spawning season, it is possible that our
collections did not coincide with the fish’s movement. Perhaps, E.
trisella were not yet present at sites sampled early in the spawning
season. Furthermore, DNA molecules may not be distributed homogenously
in water bodies due to the number of organisms upstream, river flow
rate, and the degree of water column mixing (Jerde et al., 2011).
Duplicate samples from the same localities over time would be useful in
clarifying this postulation. We plan to collect additional water samples
as well as conduct confirmatory surveys to capture live E.
trisella specimens coinciding with sites described here.
We have presented here our novel approach to noninvasive surveillance of
freshwater species. Our protocol has provided evidence that
loop-mediated isothermal amplification is a robust method for detecting
low quantities of eDNA in the water and can be modified for application
in other systems. We acknowledge that results of eDNA surveillance
efforts should be interpreted with caution and require subsequent
sequencing to confirm detection of the intended target.
ACKNOWLEDGEMENTS
This publication was made possible through a grant (62429) from the
National Fish and Wildlife Foundation (NFWF), with funding provided by
the U.S. Fish and Wildlife Service, the U.S. Forest Service, and
Southern Company. A cost share was contributed by the Geological Survey
of Alabama. The views and conclusions contained in this document are
those of the authors and should not be interpreted as representing the
opinions or policies of the U.S. Government, NFWF, or Southern Company.
Mention of trade names or commercial products does not constitute their
endorsement by the U.S. Government or NFWF or its funding sources. We
thank Kenny Jones, John Larrimore, Brianna Forrest, Anna Eastis, Autumn
Younge, and Dominque Dawson for filtering water and extracting eDNA and
R. Parker Nenstiel and Peter Dimmick for water sample collection. We
thank Brett Albanese for help with grant management.
DATA ACCESSIBILITY STATEMENT
All sequences are available on Genbank under the following accession
numbers: MT490609, MT490610, and MT490611.
AUTHOR CONTRIBUTIONS
The research project was designed by MWS with contributions from
authors; water samples were collected by AP; laboratory work and
analyses were designed and conducted by KMF and MWS; and the paper was
written by KMF with contributions from all of the authors.
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