Introduction
Originally used to assess microbial communities of ocean sediments
(Ogram, Sayler, & Barkay, 1987), the use of environmental DNA (eDNA)
applications have broadened significantly in recent decades to include
the detection and monitoring of a wide range of species in marine and
freshwater ecosystems (Martellini, Payment, & Villemur, 2005; Ficetola,
Miaud, Pompanon, & Taberlet, 2008; Jerde, Mahon, Chadderton, & Lodge,
2011; Dejean, Valentini, Miquel, & Taberlet al., 2012; Thomsen,
Kielgast, et al., 2012a). The approach has also increasingly targeted
terrestrial species using eDNA deposited within natural or artificial
water bodies (e.g. Ushio et al., 2017; Williams, Huyvaert, Vercauteren,
Davis, & Piaggio, 2018), or deposited in soils (e.g. Buxton,
Groombridge, & Griffiths, 2018; Kucherenko, Herman, III, & Urakawa,
2018; Leempoel, Hebert, & Hadly, 2019; Sales et al., 2019; Walker et
al., 2017). More recently, a number of novel techniques to collect eDNA
deposited on substrates found in aboveground terrestrial settings (e.g.
vegetation surfaces, crops, and spider webs) have broadened the
application of eDNA methods to include deployment of monitoring
protocols designed to survey terrestrial species and communities
(Nichols, Koenigsson, Danell, & Spong, 2012; Valentin, Fonseca,
Nielsen, Leskey, & Lockwood, 2018; Valentin, Fonseca, Gable, Kyle, et
al., 2020; Xu, Yen, Bowman, & Turner, 2015). However, while the state,
transport, and fate (i.e. the ‘ecology’) of eDNA in aquatic ecosystems
has been thoroughly explored (Barnes and Turner, 2016) it is not well
understood in terrestrial ecosystems, leaving key questions surrounding
sampling design and detection rates unanswered.
Understanding the state, transport, and fate of eDNA is critical to the
design and interpretation of eDNA surveys (Barnes and Turner, 2016).
eDNA can be present in multiple states; either in intracellular,
intraorganelle, or extracellular form (Turner, Barnes, Xu, Jones, et
al., 2014). Past eDNA surveys have collected a mixture of different
states through direct substrate testing (i.e. DNA extractions directly
from soil, or fecal material – Kucherenko et al., 2018; Martellini, et
al., 2005), or targeted specific states through chemical isolation (e.g.
Minamoto, Yamanaka, Takahara, Honjo, et al., 2011; Taberlet, Prud’Homme,
Campione, Roy, et al., 2012) or differential size selection via
filtration or centrifugation (e.g. Turner et al., 2014; Martellini et
al., 2005). Identifying the eDNA state(s) most common within the
environment being surveyed, or relevant to the question being addressed,
and using appropriate isolation methods to capture the desired state(s),
is key to designing protocols that maximize the probability of species
detection (Turner et al., 2014). Capture of specific eDNA states in
suspension is typically accomplished via direct processing of water,
tissue centrifugation, filtration, or a combination thereof (e.g.
Martellini, et al., 2005;
Goldberg, Pilliod, Arkle, & Waits, 2011; Jerde et al., 2011; Minamoto
et al., 2011); with filtration being the most common approach at
present. However, the existing literature guiding filtration of eDNA
states via specific filter pore sizes (e.g. Turner et al., 2014; Wilcox,
McKelvey, Young, Lowe, et al., 2015; Moushomi, Wilgar, Carvalho, Creer
et al., 2019) does not represent the full range of pore sizes that may
influence optimal capture of intracellular eDNA (i.e., trade-offs
between maximum water filtration and DNA yield). Similarly,
understanding how environmental conditions affect the decay of each eDNA
state over time informs the interpretation of positive or negative
detection results.
For instance, if a captured eDNA state persists in the environment for
long periods (i.e. months or years; e.g. Andersen, Bird, Rasmussen,
Haile, et al., 2012; Turner, Uy, & Everhart, 2015; Strickler, Fremier,
& Goldberg, 2015) it is unknown if a positive species detection
indicates a recent presence or one over a relatively long time frame.
Conversely, eDNA states that degrade quickly after deposition (i.e.
hours, days, or weeks; e.g. Zhu, 2006; Thomsen, Kielgast, Iversen, Wiuf,
et al., 2012b; Thomsen et al., 2012a) may indicate species presence
within the very recent past, or may break down beyond detectability and
produce false negative survey results (Schultz & Lance, 2015). Most
existing knowledge about the fate of eDNA comes from experiments
conducted within water or soil, finding that biotic and abiotic factors
such as pH, microbial load, temperature, and enzymatic fragmentation
influence the decay rates of eDNA (Barnes and Turner, 2016; Levy-Booth
et al., 2007; Nielsen, Johnsen, Bensasson, & Daffonchio, 2007).
However, these biotic and abiotic factors are unlikely to determine the
fate of eDNA within aboveground terrestrial systems, since eDNA in said
systems likely dries shortly after deposition and is thus likely
influenced predominantly, if not entirely, by solar radiation
(Figure 1 ).
eDNA transport in aquatic environments is facilitated by omnidirectional
diffusion, precipitation through the water column, and directional
movement via currents or thermal mixing, which can redistribute eDNA
meters to kilometers away from the point of original deposition
(Eichmiller, Bajer, & Sorensen, 2014; Deiner, Fronhofer, Mächler,
Walser, & Altermatt, 2016; Thomsen, Kielgast, et al., 2012a). These
processes can increase the availability of eDNA for capture and elevate
detection probability, or dilute the available eDNA beyond detectability
and reduce detection probability (Schultz & Lance, 2015).
Transportation of eDNA deposited within soil is not as well understood
beyond recognition that eDNA is unlikely to move laterally through the
soil substrate (Taberlet, Bonin, Coissac, & Zinger, 2018). Therefore,
the mechanisms that influence eDNA transport in water are not applicable
to aboveground terrestrial eDNA. We posit that eDNA deposited on
aboveground terrestrial surfaces will be predominately transported by
weather events like rainfall, transferring it to the soil where it may
percolate through the soil column for unknown distances (Figure
1 ). Given the unknown nature of eDNA transport in soil, surveying for
species above the soil substrate necessitates the collection of eDNA
from aboveground terrestrial substrates to ensure detection. Thus, the
use of terrestrial eDNA aggregation techniques, which pool eDNA from a
wide geographic area into a single reservoir (Valentin et al., 2018),
will become invaluable for surveys of aboveground terrestrial
environments. Aggregation of aboveground terrestrial eDNA has thus far
been executed in two ways: by directly collecting substrates and
submersing them into a centralized container filled with solution to be
sampled later (i.e. vat aggregation – Valentin et al., 2018), or
actively sampling and pooling eDNA from the substrate’s surface by
physically removing it (Valentin et al., 2020).
For collection of aboveground terrestrial eDNA en masse via
aggregation from surface substrates to move into regular use across a
variety of survey designs, including rare, threatened, or invasive
species detection and community level assessments, further investigation
of the ‘ecology’ of aboveground terrestrial eDNA is required. Here, we
conduct a series of experiments to investigate (1) the optimal filter
pore sizes for isolating extracellular aboveground terrestrial eDNA; (2)
how rain events limit eDNA retention on vegetative surfaces; and (3) the
rate of degradation due to time of air exposure and ultraviolet (UV)
solar radiation.