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\begin{document}

\title{Using eDNA to survey amphibians: Methods, applications, and challenges}



\author[1]{Xiaoxuan Sun}%
\author[1]{Ningning Guo}%
\author[1]{Jianan Gao}%
\author[1]{Nengwen Xiao}%
\affil[1]{Chinese Research Academy of Environmental Sciences}%


\vspace{-1em}



  
  \date{May 23, 2024}


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\selectlanguage{english}
\begin{abstract}
In recent years, environmental DNA (eDNA) has received attention from
biologists due to its sensitivity, convenience, labor and material
efficiency, and lack of damage to organisms. The extensive application
of eDNA has opened avenues for the monitoring and biodiversity
assessment of amphibians, which are frequently small and difficult to
observe in the field, in areas such as biodiversity survey assessment
and detection of specific, rare and endangered, or alien invasive
species. However, the accuracy of eDNA can be influenced by factors such
as ambient temperature, pH, and false positives or false negatives,
which makes eDNA an adjunctive tool rather than a replacement for
traditional surveys. This review provides a concise overview of the eDNA
method and its workflow, summarizes the differences between applying
eDNA for detecting amphibians and other organisms, reviews the research
progress in eDNA technology for amphibian monitoring, identifies factors
influencing detection efficiency, and discusses the challenges and
prospects of eDNA. It aims to serve as a reference for future research
on the application of eDNA in amphibian detection.%
\end{abstract}%



\sloppy


\section*{Introduction}

{\label{introduction}}

Amphibians are vertebrates with life cycles that crucially depend on
both aquatic and terrestrial habitats. They play an important
intermediate position in food chains and ecosystems, and are important
indicator taxa of environmental health (Li et al., 2017). Amphibians are
not only an important component of biodiversity but are also a taxonomic
group that has undergone significant changes in taxonomical organization
in recent years and is experiencing rapid population declines and risks
of extinction at a global scale (Gao et al., 2022). China is among the
countries with the richest biodiversity in the world and specifically
boasts the highest diversity of amphibian species(Xu et al.,2018).
Monitoring the status of this diverse fauna is often challenging.
According to the \emph{Technical Provisions on Survey and Assessment of
Amphibian and Reptile Diversity in County Areas} (Ministry of Ecology
and Environment Announcement No. 84, 2017), traditional methods to
investigate amphibian diversity include traditional line transect
methods, trap-barrier systems, methods using artificial covers or
shelters, mark-recapture, call counting, and others. Traditional
investigation is often accurate and intuitive, but efficiency is
frequently affected by species, life stages and environmental conditions
(Ruppert et al., 2022). Moreover, a high level of taxonomic knowledge is
required of the traditional investigator, especially for identification
of eggs and larvae (Hopkins \& Freckleton, 2002; S. Zhang et al., 2020).
In recent years, there has been increasing use of the environmental DNA
(eDNA) method, which allows the DNA of target organisms obtained from
environmental samples (e.g. water, soil, excrement, ancient sediments,
etc.) to be used for detection of presence or absence of the species,
and even enables inferences about their numbers and biomass (Ficetola et
al., 2019; Thomsen \& Willerslev, 2015). Extensive use of eDNA
technology has been made for biodiversity surveillance across diverse
taxa, encompassing fish (Deiner et al., 2017; Jerde et al., 2011; P. F.
Thomsen, J. Kielgast, L. L. Iversen, P. R. Moller, et al., 2012),
amphibians (Goldberg et al., 2018; Pilliod et al., 2013; Strickler et
al., 2015; Valentini et al., 2016), benthic fauna (Cowart et al., 2018;
Laroche et al., 2018; Stoeck et al., 2018), plants (Johnson et al.,
2019; Johnson et al., 2021; Kodama et al., 2022), bacteria (Laroche et
al., 2018; Stoeck et al., 2018; Y. Zhang et al., 2020), and
microorganisms (Handelsman, 2004; Rondon et al., 2000). Nonetheless,
varied applications of eDNA technology among distinct biological cohorts
elicit discernible disparities. This review provides an exposition of
the applications of eDNA technology in amphibian monitoring and also
discusses the multifaceted factors determining its efficacy and
reliability. We also examine future research and questions that will
need to be addressed for the further development of eDNA technology for
amphibian population investigations.

\section*{1 Introduction of eDNA
method}

{\label{introduction-of-edna-method}}

Due to the intensification of human activities and increased
interference with the natural environment, the continuous decline of
global biodiversity has become one of the major challenges in the 21st
century (Butchart et al., 2010). Rapid, sensitive and accurate
biodiversity survey tools are required to measure fluctuations in
species diversity and abundance to inform conservation and management
strategies (Kelly et al., 2014). Environmental DNA is DNA that can be
extracted from environmental samples (such as soil, water or air)
without prior isolation of any target organisms (Lodge et al., 2012;
Taberlet et al., 2012). Environmental DNA analysis is divided into two
main categories: single species identification and eDNA metabarcoding.
Single species identification is mainly used for sensitive detection of
rare and endangered animals and invasive non-native species in the
context of environmental and species richness surveys (Wilcox et al.,
2013). eDNA metabarcoding refers to analysis of eDNA with
high-throughput sequencing technology, processing eDNA from water, soil
and air by PCR amplification to determine the presence of species and
perform biodiversity assessment (Ruppert et al., 2019). Compared with
traditional survey methods, the eDNA metabarcoding technique provides
outstanding species recognition. It can be used even if the
investigators lack extensive abilities to classify species. Further, the
use of eDNA does not require sampling approaches that are destructive or
invasive to the environment or the biological community, nor does it
require special equipment or particular observation times to detect
those species that are difficult to observe due to small size or numbers
(Deiner et al., 2017). Therefore, this technology has been widely used
in the field of ecology.

\section*{2 Key aspects and challenges in eDNA monitoring of
amphibians}

{\label{key-aspects-and-challenges-in-edna-monitoring-of-amphibians}}

\subsection*{2.1 Sample collection and DNA
capture}

{\label{sample-collection-and-dna-capture}}

Amphibians exhibit a bifurcated life history, comprising both aquatic
and terrestrial phases. In the former, their eDNA is disseminated into
the aquatic milieu via exudates from integumentary secretions, excreta,
and ova deposition. In the latter, eDNA is presumed to derive from
imprints generated during their terrestrial locomotion, manifesting as
soil or sedimentary remnants. Consequently, a comprehensive scrutiny of
amphibian eDNA mandates the simultaneous procurement of aquatic and
terrestrial substrates. The sources of collecting amphibian
environmental DNA samples are mainly water, soil, and other natural
substrates. Some studies have demonstrated that detecting terrestrial
vertebrates from soil samples is challenging and that eDNA testing is
more complex under these circumstances. For example, Walker et al.
cultured salamanders in a mesocosm ecosystem and found that only 1\% of
salamander DNA from soil samples could be successfully amplified (Walker
et al., 2017). The success of applying the eDNA method for detection
depends not only on the presence and concentration of eDNA in the
substrate sample but also on capture potency, DNA extraction success,
sample interference (such as inhibition), and assay sensitivity
(Ficetola et al., 2015; MacKenzie et al., 2002; Schultz \& Lance, 2015).
Filtration is the standard approach for the capture of DNA from aquatic
environments (Goldberg et al., 2016); generally, volumes between 15 mL
and 10 L of water are sampled to trap eDNA in a filter. In contrast to
fish, amphibians may exhibit comparatively modest eDNA exudation. Some
judicious augmentation of filtrate volume may therefore be considered to
enhance the sensitivity of detection. Common filtration materials
include nitrate cellulose (Caren S Goldberg et al., 2011), glass fiber
(Jerde et al., 2011), polycarbonate (Takahara et al., 2012), nylon
(Philip Francis Thomsen et al., 2012) and polyether sulfone (Renshaw et
al., 2015). Chen et al. evaluated the ability of 12 different types of
adsorption materials and filters to capture eDNA in both laboratory and
field experiments using quantitative PCR (qPCR) analysis. They found
that the ability of a glass fiber filter (GF) to capture eDNA was
superior to other materials, and that the amount of eDNA captured
increased linearly over 72 hours (Chen et al., 2022).

It has been demonstrated that the period of detectability of eDNA in
water samples is limited to between 1 day and 8 weeks (Dejean et al.,
2011; Pilliod et al., 2014; Philip Francis Thomsen et al., 2012; P. F.
Thomsen, J. Kielgast, L. L. Iversen, C. Wiuf, et al., 2012), and
therefore false negative results may be obtained if there is a delay in
the collection of water samples relative to the time of species
presence. In addition, temperature alterations caused by seasonal
changes will also affect the content of eDNA in samples. For example,
Wei et al. studied the variation over time of eDNA in sediments over
periods of one year, one month and one tidal period, and found that eDNA
content was higher in the warm season and lower in the cold season (Wei
et al., 2019). Similarly, the concentration and distribution of eDNA can
be influenced by spatial factors such as spatial dissipation of eDNA and
the flow of the water column. In still water, the eDNA of species
decreases significantly within a radius of 5--10 m from the eDNA point
source (Brys et al., 2021), while in flowing waters, eDNA may move with
the current to new locations hundreds of meters away (Pilliod et al.,
2013), leading to false negatives at the original point as well as false
positives at the new location.

Therefore, biases in results due to temporal and spatial differences
need to be taken into account when designing eDNA experiments, and these
factors also need to be considered when conclusions are drawn about the
range of species present. For example, conclusions drawn from water
samples taken along one side of a river are not directly representative
of the whole river.

\subsection*{2.2 DNA extraction}

{\label{dna-extraction}}

During amphibian DNA extraction, pollution should be avoided and the
facilities and equipment employed should be disinfected. Standard
autoclave methods are not sufficient to destroy nucleic acid
molecules(Unnithan et al., 2014). Among common stain removal methods,
sodium hypochlorite solvent is the most effective for removing DNA and
PCR products (Champlot et al., 2010; Prince \& Andrus, 1992). The choice
of eDNA extraction method after filtration can greatly affect the
detection of eDNA. Deiner et al. compared the effectivity of six
different combinations of sample collection and DNA extraction methods
in biodiversity monitoring in water ecosystems. The results showed that
different environmental DNA collection and extraction methods had a
significant effect on DNA yield and the number of sequences obtained by
high-throughput sequencing techniques (Deiner et al., 2015).

Common DNA extraction methods include liquid phase separation and
commercial DNA extraction kits. Trials have shown that liquid phase
separation methods can in some cases outperform kits in terms of
extraction efficiency. Turner et al. showed that a
cetyltrimethylammonium bromide (CTAB) method using polycarbonate orbital
filters collected more eDNA than a PowerWater DNA isolation kit using GF
membranes (1.5 \selectlanguage{greek}µ\selectlanguage{english}m pore size, 934-AH grade) (Turner et al., 2014). It has
also been shown that the phenol-chloroform-isoamyl alcohol extraction
(PCI) method using glass and nitrocellulose membranes extracted more DNA
than the DNeasy kit (Deiner et al., 2015; Renshaw et al., 2015).
However, liquid phase separation methods are difficult to apply widely
due to the utilization of toxic substances such as phenol and chloroform
(Tsuji et al., 2019).

\subsection*{2.3 Primer selection and PCR
amplification}

{\label{primer-selection-and-pcr-amplification}}

Species-specific detection is carried out the basis of PCR techniques
that use specific primers to amplify and detect short segments of DNA
(typically 80--200 bp) of the target species (Bohmann et al., 2014).
eDNA was first applied to large organisms using specific primers that
amplified only one species during PCR (Jerde et al., 2011; P. F.
Thomsen, J. Kielgast, L. L. Iversen, C. Wiuf, et al., 2012). Advances in
sequencing technology have made it possible to assess the community
composition using universal primers that identify multiple species in a
sample; universal primers are required to amplify all species in a
target taxonomic unit with a minimal bias to avoid preferential
amplification of some species with mismatched primer regions (Ficetola
et al., 2010). The mitochondrial genes \emph{12S} and \emph{Cytochrome
b} and the mitochondrion's D-loop region are commonly used in amphibian
studies, and the \emph{12S} region is often used for metabarcoding eDNA
analysis (Table 1) (Li et al., 2021; Wynne E Moss et al., 2022; Sasso et
al., 2017). For example, Valentini et al. designed a pair of primers
specific for amphibians and scleractinians (batra\_F,
5'-ACACCGCCCGTCACCCT-3,' batra\_R, 5'-GTAYACTTACCATGTTACGACTT-3') and a
human DNA blocking primer (batra\_blk, 5'-
TCACCCTCCTCAAGTATACTTCA-AAGGCA-SPC3I-3'); this primer pair can amplify
up to 99\% of amphibian species (Valentini et al., 2016). In a previous
study, we employed GF filtration membranes in conjunction with the
``batra'' primers to investigate the amphibian diversity within China's
Chao Lake(Unpublished data). This approach not only facilitated the
successful amplification of all species identified through traditional
sampling methods but also unveiled two species that had previously
eluded detection in historical records and conventional surveys. These
outcomes substantiate the efficacy of the 'batra' primers for detecting
Chinese amphibians.

Because it is impossible to pinpoint organisms directly based on eDNA
molecules exuded into water, nor to track the movement of specific eDNA
molecules in water, the environmental samples collected often contain
mixed DNA from many different organisms. When amplifying environmental
samples, the specific primers chosen often play an important role in the
successful amplification of the target species. Unsuitable primers have
the potential to generate false positives, which can lead to
overestimates of the presence or abundance of endangered species, or to
false negatives, which can lead to insufficient monitoring of invasive
species. In his study, Kelly pointed out that using a constant number of
PCR cycles, different primers showed different biodiversity structures
for the same simulated community (Kelly et al., 2019), which directly
led to different analysis results. Wilcox tested the factors affecting
the specificity and sensitivity of eDNA detection, and found that
specificity was most affected by the mismatch of base pairs in the
primers, which was much more prevalent than in the probe (Wilcox et al.,
2013). Hence, it is evident that the meticulous selection of appropriate
primers is of paramount importance.

\begin{equation}
Cq=(\text{Cq}_{\text{samples}}-\text{Cq}_{\text{positive\ control}})\nonumber \\
 \end{equation}

It has been demonstrated that diluting the sample alleviates the
inhibition, but likewise dilutes the target DNA. Especially when the
target DNA is present at very low concentrations, this may lead to
failure of the assay (Goldberg et al., 2013; Anna M McKee et al., 2015;
Strand et al., 2011; Tsai \& Olson, 1992). McKee et al. examined this
using a PCR inhibitor removal kit and found some alleviation of
inhibition but also potential loss of target DNA (Anna M McKee et al.,
2015).

\subsection*{2.4 Sequencing and data
analysis}

{\label{sequencing-and-data-analysis}}

The PCR results are recorded as negative or positive. A negative result
indicates that eDNA of the target species is not present in the
environmental sample, while a positive result indicates its presence.
This result is initially determined by GEP after primer amplification
and then further verified by sequence comparison. In qPCR, a
fluorescence threshold is set using the positive control as a benchmark;
the sample is labelled positive when the amplification exceeds the
fluorescence threshold, and negative when it does not (Venkatesan \&
Bashir, 2011). Metabarcoding uses universal PCR primers to amplify DNA
in the environment. A large amount of data containing taxonomic
information on genes can be obtained after high-throughput sequencing,
which requires sequence comparison and species annotation (including
noise removal and assignment of operational taxonomic units),
clustering, and assignment of species annotation.

A complete and high quality reference database is the basis for the
accuracy of amphibian surveys using eDNA technology (Alexander et al.,
2020; West et al., 2022). This can be constructed by consulting
established publicly available databases (e.g. NCBI:
https://www.ncbi.nlm.nih.gov/) or by building custom versions. For
example, Valentini et al. built a reference database containing 53
species of amphibians when using eDNA method to investigate amphibian
diversity (Valentini et al., 2016). The accuracy of self-built databases
is likely to be higher than that of public databases due to the
significant geographic variation in amphibian fauna and the lack of
assurance of the quality of DNA barcoding in publicly available
databases.

\section*{3 Progress of domestic and international eDNA-based surveys of
amphibians}

{\label{progress-of-domestic-and-international-edna-based-surveys-of-amphibians}}

\subsection*{3.1 Species diversity
surveys}

{\label{species-diversity-surveys}}

With the signing of the Convention on Biological Diversity (CBD) and the
15th Conference of the Parties (COP15) to the United Nations Framework
Convention on Climate Change (UNFCCC), global biodiversity has received
wide attention. eDNA technology combined with Next-generation sequencing
offers a more sensitive and efficient method for biodiversity surveys.
Svenningsen et al. used simultaneously traditional survey methods and
eDNA metabarcoding to detect amphibian diversity in Danish lakes and
ponds, and evaluated the detection efficiency of the two methods from
the perspectives of species richness, the average number of species
detected at various points, and frequency of detection. They found that
both eDNA metabarcoding and traditional surveys yielded a large number
of observations unique to the respective method, suggesting that eDNA
metabarcoding should be combined with traditional methods as a
complement to enhance determination of the total composition of species
(Svenningsen et al., 2022).

eDNA detection of amphibians has yielded good results in temperate
regions (Deiner et al., 2017; Thomsen \& Willerslev, 2015; Yates et al.,
2019). However, it has been demonstrated that high temperatures and
strong UV light favor the growth of microorganisms and promote the
degradation of DNA. To validate eDNA metabarcoding surveys of amphibian
diversity in the tropics, Sasso et al. analyzed four streams in the
Atlantic Forest of southeastern Brazil during a five-year period of
traditional field surveys (using visual and acoustic methods) as well as
short-term sampling (4 days) of eDNA. A total of 10 species of aquatic
amphibians were detected over the 5-year period of the traditional
survey, while nine species were detected by eDNA metabarcoding,
including one species that had only been detected once during the 5-year
traditional survey. There was a high degree of similarity in stream
community composition between the results of the two methods (Sasso et
al., 2017). Li et al. used eDNA metabarcoding to assess amphibian
diversity at 288 sites in 18 regions of Hainan Island and found 15
species, including a number difficult to detect with traditional
monitoring methods (Li et al., 2021). These studies demonstrate the
effectiveness of the eDNA metabarcoding technique in detecting
amphibians in the tropics.

\subsection*{3.2 Detection of invasive
species}

{\label{detection-of-invasive-species}}

Alien invasive species are one of the main causes of global biodiversity
loss and homogenization (Ehrenfeld, 2010; Ficetola et al., 2007; Pysek
\& Richardson, 2010; Vitousek et al., 1997). Invasive species may
disrupt local ecological balance and disturb the ecosystem, with
substantial impacts on the variety and abundance of native species
(Ficetola et al., 2008). The early presence of invasive species is
difficult to detect in a timely manner using traditional methods, while
eDNA technology allows for rapid and sensitive monitoring of invasive
organisms. Invasive amphibians were the first vertebrates for which
species distribution was successfully assessed by extracting DNA from
water samples (Ficetola et al., 2008). Specific primers were used to
expand short mitochondrial DNA sequences to trace the presence of
invasive bullfrogs (\emph{Rana catesbeiana} ) in controlled environments
and natural wetlands. Similarly, Dejean et al. assessed the distribution
of this species in ponds in western France using traditional monitoring
(visual and acoustic surveys) and eDNA. Bullfrogs were found in only
14\% of sites in the traditional survey, while the eDNA survey detected
a much higher presence (78\%), demonstrating substantial effectiveness
and sensitivity (Dejean et al., 2012).

\subsection*{3.3 Detection of rare and endangered
species}

{\label{detection-of-rare-and-endangered-species}}

eDNA has great potential for detecting rare and endangered species.
Pierson et al. investigated the occurrence of the tiny, rare,
multi-toothed salamander \emph{Urspelerpes brucei} by using both the
litter bag survey and eDNA methods. The results showed that the
detection probability of the eDNA survey was 0.788, much higher than
that of the litter bag survey (0.048). The eDNA method also imposes
weaker disturbances on aquatic habitats and has lower costs than the
litter bag method (Pierson et al., 2016). Goldberg et al. designed
specific primers to detect two rare species in the United
States,\emph{Ascaphus montanus} and \emph{Dicamptodon aterrimus} . In
the experiment, water samples were collected from five rivers with
different densities of the target species, and PCR amplification was
performed to detect whether the species' DNA existed in the
samples.\emph{Dicamptodon} was detected in all water samples and
\emph{Ascaphus}in four of the five rivers, proving the effectiveness of
eDNA detection for surveys of these rare species (Caren S Goldberg et
al., 2011).

McKee et al. studied the species \emph{Ambystoma cingulatum}
,\emph{Ambystoma bishopi} , \emph{Notophthalmus perstriatus}
and\emph{Lithobates capito} in the southeastern longleaf pine region of
the United States using both eDNA and traditional methods (mainly
dip-netting, trapping, and visualization). The eDNA method succeeded in
detecting the target species at six sites where traditional methods did
not show a presence, again indicating that eDNA can be used as a
supplement to traditional methods for investigating endangered species
(A. M. McKee et al., 2015). Voros et al. used eDNA to detect the
presence of the rare cave-dwelling amphibian \emph{Proteus anguinus} in
all of 15 sampled caves in Croatia, five of which constituted first
records for the species (Voros et al., 2017). Lopes et al. analyzed
water samples from the Atlantic Coast Forest and six mountainous areas
of the adjacent Cerrado grasslands in Brazil for traces of DNA of rare
and endangered amphibian species. They successfully detected four
declining species (\emph{Hylodes ornatus} , \emph{Hylodes regius}
,\emph{Crossodactylus timbuhy} and \emph{Vitreorana eurygnatha} ), two
species considered locally extirpated (\emph{Phasmahyla exilis}
and\emph{Phasmahyla guttata} ), and a species not seen since 1968
(\emph{Megaelosia bocainensis} ) (Lopes et al., 2021). These studies
successfully confirmed the presence of species not detected by
traditional methods, and underline the effectiveness of the eDNA method
for biodiversity monitoring at low population densities.

\subsection*{3.4 Species abundance and biomass
assessment}

{\label{species-abundance-and-biomass-assessment}}

eDNA allows for the assessment of the relative abundance and biomass of
faunal communities. In 2019, Kelly et al. analyzed eDNA macro-barcodes
of three simulated biomes with different biomass ratios. The effects of
these ratios on biodiversity estimates were analyzed by assuming that
the amount of DNA collected in the environment was proportional to
abundance of individuals in the water, and that the DNA shedding rate
and amplification efficiency of certain primers of isotaxa was constant.
The analysis results showed that in the case of high amplification
efficiency, such as primer adaptation, the ratio index of eDNA
macro-barcode readings was highly indicative of trends in group biomass
(Kelly et al., 2019). This result has been replicated in amphibian
studies. For example, Thomsen et al. investigated six different species
including two amphibians (\emph{Pelobates fuscus} and \emph{Triturus
cristatus} ) using eDNA and qPCR methods, finding that the eDNA
concentrations of these two species in water samples correlated with
density and biomass (P. F. Thomsen, J. Kielgast, L. L. Iversen, C. Wiuf,
et al., 2012). Pilliod et al. used eDNA and traditional methods to
assess the diversity and abundance of two amphibian
species,\emph{Ascaphus montanus} and \emph{Dicamptodon aterrimus} , in
13 streams in central Idaho, USA. They reported that eDNA reading
proportion was positively correlated with density and biomass as
measured by traditional methods, and showed that the accuracy of
eDNA-based abundance assessments increased with the amount of eDNA in
water and the number of replicate samples collected (Pilliod et al.,
2014). Everts et al. used droplet digital PCR (ddPCR) analysis of eDNA
to detect the abundance of bullfrogs in the United States, and found
that eDNA concentration increased significantly with increasing
abundance of bullfrog eggs and tadpoles (Teun Everts et al., 2021).

\subsection*{3.5 Reconstruction of ancient
ecosystems}

{\label{reconstruction-of-ancient-ecosystems}}

eDNA for amphibian surveys is mainly used to detect the presence of
target species in current communities. However, it has been shown that
since DNA molecules can exist in the environment for a long time,
ancient ecosystems can be assessed by extracting remnant DNA molecules
from environmental sources. Ficetola and Taberlet detected eDNA of
several amphibian species (\emph{Bufo bufo} , \emph{Rana temporaria}
and\emph{Ichthyosaura alpestris} ) in 1,000-year-old sediments from a
lake in the French Alps. This suggests that eDNA stored in lake
sediments can be used to understand changes in species distribution over
time. In addition, it can be combined with other paleoecological data to
understand species' responses to environmental changes (such as habitat
alterations, climate change, and the introduction of alien species)
(Giguet-Covex et al., 2019).

\subsubsection*{3.6 Amphibian disease
surveillance}

{\label{amphibian-disease-surveillance}}

In addition to factors such as habitat degradation and environmental
pollution, viral infections have also emerged as a significant
contributor to the decline of amphibian populations (Rachowicz et al.,
2006). According to Scheele et al., the amphibian chytrid fungus
(\emph{Batrachochytrium dendrobatidis} ; Bd) has led to the decline of
at least 501 amphibian species worldwide, with 90 species having gone
extinct, including the Darwin's frog (\emph{Rhinoderma darwinii} )
(Scheele et al., 2019). In recent years, eDNA methods have demonstrated
remarkable capabilities for detecting Bd due to their convenience,
sensitivity, and non-invasive nature.

Researchers like Stephen et al. have successfully determined the
presence and quantity of chytrid pathogens by collecting skin swabs from
amphibian hosts, extracting DNA from these swabs, and employing qPCR
techniques (Boyle et al., 2007). Beyond skin swab collection, pathogenic
DNA can also be detected from environmental samples. Walker et al., for
instance, successfully detected the presence of the Bd in small-volume
(\textless{}1 L) water samples by filtering and utilizing qPCR (Walker
et al., 2007). Similarly, Julie et al. employed filtration to capture
zoospores from water samples and subsequently detected pathogen DNA from
filtered particles at concentrations as low as 0.06 zoospores per water
sample, highlighting the high analytical sensitivity of the eDNA method
(Kirshtein et al., 2007).

The potential of eDNA technology for early detection of Bd in the
environment is noteworthy. Colleen et al. collected eDNA from filtered
water samples and detected Bd at three sites, all of which a month later
turned out to experience Bd-induced mortality, while Bd was not detected
in sites without later mortality (Kamoroff \& Goldberg, 2017). This
underscores the potential of eDNA technology for early detection of Bd
presence.

\section*{4 Influential factors of eDNA
detection}

{\label{influential-factors-of-edna-detection}}

Lindahl showed that the detection and quantification of DNA in specific
freshwater animal species depends on DNA release and degradation rates.
eDNA release is related to population density and organism size, and DNA
degradation is determined by a simple relationship between microbial or
enzymatic attack and spontaneous chemical reactions such as hydrolysis
and oxidation (Lindahl, 1993). Conceptually, three processes determine
the detection of DNA in environmental samples: production, dissipation
and degradation.

\subsection*{4.1 Production of eDNA}

{\label{production-of-edna}}

For aquatic eDNA, production, the rate at which DNA is released in
water, is a highly variable function, influenced by population density
and species-specific characteristics, such as individual size and
metabolic rate (Strickler et al., 2015). Thomsen conducted Pearson
correlation analysis of population size and the mean number of DNA
molecules in ponds for two amphibian species (\emph{Pelobates fuscus}and
\emph{Triturus cristatus} ) and found that DNA release increased with
population density and animal size (P. F. Thomsen, J. Kielgast, L. L.
Iversen, C. Wiuf, et al., 2012). In order to examine the effect of
different growth stages of amphibians on the efficiency of eDNA
detection, Everts et al. compared the eDNA emission rates of tadpoles
and juveniles of American bullfrogs (\emph{Rana catesbeiana} ) and found
that the average individual eDNA emission rate of tadpoles was not
significantly different from that of juveniles, while the average
emission rate per unit biomass of tadpoles was significantly lower than
that of juveniles (Teun Everts et al., 2021).

\subsection*{4.2 Dissipation of eDNA}

{\label{dissipation-of-edna}}

The dissipation pattern of eDNA was shown to vary between species and
experimental conditions. In order to obtain more accurate results for
amphibian detection by eDNA methods, Brys et al. estimated the temporal
and spatial dissipation of eDNA by placing a group of different fish and
amphibians in a cage on one side of a pond (thereby creating eDNA
emission sites) and assessing the dissipation of eDNA with distance by
collecting water samples at different distances from the cage. Detection
rate of the locally released eDNA and the standardized index of relative
eDNA abundance of the cage species decreased significantly with distance
within a radius of 5--10 m from the cage. After removing the cage
community for one week, no DNA of any species could be detected in the
study system, indicating that the eDNA had degraded below the detection
threshold within that period (Brys et al., 2021). Dejean et al.'s study
found that eDNA from American bullfrog (\emph{Rana catesbeiana} )
tadpoles was able to persist for 25 days under a laboratory microscope
(Dejean et al., 2011). Thomsen et al. found that in experiments
conducted in outdoor ponds, eDNA from larvae of the
amphibians\emph{Pelobates fuscus} and \emph{Triturus cristatus} could be
detected for 7--14 days (P. F. Thomsen, J. Kielgast, L. L. Iversen, C.
Wiuf, et al., 2012).

\subsection*{4.3 Degradation of eDNA}

{\label{degradation-of-edna}}

Studies have shown that degradation of DNA in water is one of the main
factors that reduce detectability (Barnes et al., 2014; Dejean et al.,
2012; Pilliod et al., 2014). An understanding of the relevant
environmental factors is therefore essential for controlling degradation
rate and optimizing sampling strategy when using eDNA methods. DNA in
water is broken down by chemical hydrolysis, primarily by exposure to
acids or by enzymatic action. Microbial activities in water directly
promote enzymatic hydrolysis by producing exogenous nucleases, which
break down DNA into its components (Lindahl, 1993).

High temperatures accelerate the degradation of DNA (Corinaldesi et al.,
2008; Okabe \& Shimazu, 2007). When temperatures are very high
(\textgreater{}50\selectlanguage{ngerman}°C), direct denaturation can occur; however, most of
the effects of high temperatures on DNA degradation arise indirectly, as
moderately high temperatures stimulate microbial metabolism and
exonuclease activity (Corinaldesi et al., 2008; Fu et al., 2012;
Hofreiter et al., 2001; Pote et al., 2009; Zhu, 2006). Strong
ultraviolet radiation can also enhance DNA degradation. This
particularly applies to ultraviolet B (UV-B) light, as UV-B wavelengths
are the most harmful to aquatic organisms among wavelengths most likely
to occur in terrestrial sunlight (Diffey, 2002), causing photochemical
damage to DNA and thus inhibiting DNA amplification (Hader et al., 2003;
Ravanat et al., 2001). Acidic conditions catalyze the hydrolysis process
that degrades DNA (Alaeddini et al., 2010; Lindahl, 1993), and thus eDNA
is present for longer in samples with neutral or slightly alkaline pH
(Lindahl, 1993).

A study by Pilliod et al. clearly quantified the influence of specific
environmental factors on eDNA degradation. eDNA of \emph{Dicamptodon
aterrimus} can persist for 8 days in an outdoor container under ambient
light and temperature conditions, at least 11 days under ambient
temperature and low light conditions, and for at least 18 days under
refrigeration without light (Pilliod et al., 2014). Similarly, to
quantify the independent and interactive effects of environmental
factors on eDNA persistence, Strickler et al. measured eDNA degradation
rates at different levels of UV-B, pH, and temperature in a laboratory
setting. The results showed that temperature had a strong effect on the
amount of eDNA detected over time; the degradation rate at 5°C was
significantly lower than that at 20°C and 35°C, indicating slower
microbial growth at low temperatures, which may delay the microbial
degradation process of eDNA. There was no significant difference in
degradation rate between pH 4 and pH 7, but rates at both pH levels were
higher than that at pH 10. There was no significant difference in the
degradation rate of UV-B at 2, 25 and 50 KJ/m\textsuperscript{2}/day,
however, the degradation rate was faster at higher UV-B levels when the
level was between 5 and 25 KJ/m\textsuperscript{2}/day and the amount of
residual eDNA was lower than 5\%. This indicates that there is a
nonlinear correlation between UV-B intensity and eDNA degradation
(Strickler et al., 2015). Since geographical features such as latitude
and altitude are the main determining factors of UV-B radiation (Godar,
2005), radiation effect on eDNA degradation is more significant in
aquatic environments at high altitudes or near the equator. Although
studies have shown that higher acidity (pH 4) accelerates the
degradation of eDNA, McKee et al. successfully detected four endangered
amphibian species (\emph{Ambystoma cingulatum} , \emph{Ambystoma
bishopi} , \emph{Notophthalmus perstriatus} and \emph{Lithobates capito}
) in natural acidic wetlands (pH close to 4) of longleaf pine in the
southeastern United States (A. M. McKee et al., 2015). Goldberg et al.
also detected the endangered species \emph{Ambystoma bishopi}
and\emph{Pseudacris ornata} at the Eglin Air Force Base and demonstrated
that a pH value as low as 4 would not degrade eDNA to an undetectable
state (Goldberg et al., 2018).

\section*{5 Prospects}

{\label{prospects}}

The implementation of eDNA methodologies for amphibian surveillance
encompasses a sequence of five primary stages: sample acquisition, DNA
extraction, PCR amplification, sequencing, and data analysis (Figure 1).
It is evident from this review that the heterogeneous array of research
objectives often engenders considerable divergence within the procedural
framework. Noteworthy variables encompass the volumetric scale of
aquatic samples, ranging from 15 mL to 10 L, the selection of filter
pore sizes spanning the interval from 0.22 \selectlanguage{greek}μ\selectlanguage{english}m to 1 \selectlanguage{greek}μ\selectlanguage{english}m, and the
deliberate designation of genetic loci such as 12S, 16S, and Cytochrome
b, each of which shows pronounced heterogeneity. This intricate
landscape precludes the facile derivation of a universally 'omnipotent'
experimental protocol. It should be noted that attempts have been made
to propagate standardized methodologies for eDNA techniques (Bruce et
al., 2021; De Brauwer et al., 2022; Minamoto et al., 2021), however
these so far have not encountered widespread acceptance or practical
implementation (Takahashi et al., 2023).

The multifaceted paradigm underlying eDNA application, as discussed in
section 2 of this review, illustrates the causes for the limited
applicability of 'standardization guidelines' across diverse contexts.
The unique physicochemical characteristics of different environments
make it necessary to carry out iterative adjustments to the experimental
approach blueprint and the methodology, as suitable to varied research
goals (Pawlowski, Apoth\selectlanguage{ngerman}éloz-Perret-Gentil, et al., 2020; Pawlowski,
Apothéloz\selectlanguage{english}-Perret\selectlanguage{english}-Gentil, et al., 2020; Taberlet et al., 2018; Takahashi
et al., 2023).

Because spatial and temporal differences have a large and unavoidable
effect on eDNA capture, it may be possible to improve the accuracy of
eDNA methods for investigating species diversity or richness of plants
and animals by combining them with distribution or occupancy models.

A high-quality and accurate reference database is required for eDNA
metabarcoding, and the information base must be sufficient to cover all
species in that experimental region. Errors or gaps in the database will
lead to a decrease in the accuracy of the findings (Abad et al., 2016;
\selectlanguage{polish}Š\selectlanguage{english}igut et al., 2017; Yang et al., 2017). Continued enrichment and
improvement of DNA barcode databases in the future are therefore
desirable to improve their accuracy and credibility.

Despite the convenience, accuracy and low cost of the eDNA method, it
has many disadvantages compared to traditional surveys, such as the
inability to directly observe the life stages and disease conditions of
surveyed organisms or to measure organism indicators. eDNA metabarcoding
cannot completely replace traditional survey methods, and choice of
survey methodology should be weighed against the advantages and
limitations of both types of approach, as well as specific research
objectives. Complementary use is most likely to achieve the most
desirable research results.

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\section*{Tables}

{\label{tables}}

\textbf{Table1.} Primers used in eDNA amphibian surveys

\textbf{Aim}

\textbf{Research Area}

\textbf{Mitochondrial Genome Region}

\textbf{Primer Name}

\textbf{Forward Primer(5'-3')}

Reverse Primer(5'-3')

\textbf{Length}

\textbf{Reference}

Threatened Species (\emph{Latonia nigriventer})

Northern Israel

12S

/

(5'- GAACTACGAGCCTCAGCTTAAA-3') (5'- GGCAAGAAGTGGTGAGGTTA-3')

110 bp

(Renan et al., 2017)

Threatened Species (\emph{Ambystoma cingulatum}, \emph{Ambystoma
bishopi})

Southeastern United States

D-loop

Amcin F Amcin R Amcin P

(5'-GGCCCGTCAACTTTCCTCTAA-3') (5'-TGGTCCAGGTAAATCAATTGCA-3')
(6FAM-TACGGTAATATGTCTGGTACTAC-MGBNFQ)

132 bp

(Anna M McKee et al., 2015)

Threatened Species (\emph{Notophthalmus perstriatus})

NADH Dehydrogenase Subunit 1 (ND1)

Noper F Noper R Noper P

(5'-CCGAGCCCCCTTCGAT-3') (5'-CTTCCTGCATATTCTACGTTAAATCCT-3')
(6FAM-TAACAGAAGGTGAATCTG-MGBNFQ)

71 bp

Threatened Species (\emph{Lithobates capito})

NADH Dehydrogenase Subunit 2 (ND2)

Licap F Licap R Licap P

(5'-CGGCACTACAGTCACCCTATC-3') (5'-TCGGGATAATGGCGAGGGTAT-3') (CAL Fluor
Red610-TYCATTGACTCTTAGCCTGAGTAGGCYTA-BHQ-2)

82 bp

Threatened Species (\emph{Phytotriades auratus})

Trinidad, America

Cytb

GTF-F GTF-R

(5'-CCCCTT ACATCGGCACTGAC-3') (5'-CTCCAAGGATGTTTGGGGTGA-3')

390bp

(Brozio et al., 2017)

Threatened Species (\emph{Babina subaspera})

Amami-Oshima Island,Japan

ND5

BsubND5-F BsubND5-R

(5'-TCACTCAATGCCCGCTTATATA-3') (5'-GTACTCTAAAACCAATTTAGTA-3')

90 bp

(Takahara et al., 2020)

Threatened Species (\emph{Odorrana splendida})

ND5

OsplND5-F OsplND5-R

(5'-GCTGAGAAGGCGTAGGATTT-3') (5'-ACATCTCATGTCGAACATTGC-3')

96 bp

Threatened Species (\emph{Odorrana amamiensis})

ND5

OamaND5-F OamaND5-R

(5'-TCGGCCCAGGCCCTTTTGAGCA-3') (5'-TGATTGTGTTCCCACTA-3')

116 bp

Rare Species (\emph{Proteus anguinus})

Dinaric Karst, Croatia

D-loop

Paf8 Par8

(5'-GTGGCATATAAATCTATGTC-3') (5'-TRTTATTCGTTTTCTAGG-3')

64 bp

(V\selectlanguage{ngerman}örös et al., 2017)

Rare Species (\emph{Phytotriades auratus})

West Indies

Cytb

/

(5'- GCGGATTCTCTGTCGACAATG -3') (5'- TGCTCCTGCAATAAGAAATGGA -3')

80bp

(Torresdal et al., 2017)

Secretive Species (\emph{Ascaphus montanus})

Northwest United States

Cytb

Asmo F Asmo R

(5'-CGTCAACTATGGCTGGCTAA-3') (5'-TCGGCCAATGTGAAGATAAA-3')

78 bp

(C. S. Goldberg et al., 2011)

Secretive Species (\emph{Dicamptodon aterrimus})

Dicamp F Dicamp R

(5'-TCTGCATCTTYCTACATATYGGAC-3') (5'-ATCACYCCGACKTTTCAGGT-3')

85 bp

Alien Invasive Species Quantitative Detection (\emph{Lithobates
catesbeianus})

Western European

Cytb

BullfrogF BullfrogR

(5'-TTTTCACTTCATCCTCCCGTTT-3') (5'-GGGTTGGATGAGCCAGTTTG-3')

84 bp

(T. Everts et al., 2021)

Alien Invasive Species Quantitative Detection (\emph{Lithobates
catesbeianus})

Western European Beijing, China

16S

qLC16S Forward qLC16S Reverse

(5'-GCAGAGATAACCTCTCGT-3') (5'-GTCCCATAGGACTGTTCT-3')

120 bp

(T. Everts et al., 2021) (Lin et al., 2019)

Alien Invasive Species (\emph{Lithobates catesbeianus})

Southwest of France

Cytb

/

(5'-TGCCAACGGAGCATCATTC-3') (5'-ATAAAGGTAGGAGCCGTAGT-3')

79 bp

(Dejean et al., 2012)

Quantitative Detection

Linkebeek, Belgium

12S

12S\_F1 12S\_R1

(5'-ACTGGGATTAGATACCCC-3') (5'-TAGAACAGGCTCCTCTAG-3')

142 bp

(Brys et al., 2021)

Quantitative Detection (\emph{Ascaphus montanus})

Idaho, United States

Cytb

qASMOF qASMOR

(5'-ACGTCAACTATGGCTGGCTAATC-3') (5'-GTCCTCGGCCAATGTGAAGA-3')

within 200 bp

(Pilliod et al., 2013)

Quantitative Detection (\emph{Dicamptodon aterrimus})

qDIATF qDIATR

(5'-AAACTTTACACCAGCAAACCCATT-3') (5'-CGGAGAATAGCGTATGCAAAAA-3')

Species-specific detection (\emph{Hynobius kimurae})

Kyoto Prefecture, Japan

12S

Hynobius\_12S\_F1 Hynobius\_12S\_R1 Hida\_12S\_P

(5'-TTAATAAAAACGGCCTAAAGCGTG-3') (5'-TCAATTATAGAACAGGCTCCTCTAGGG-3')
({[}FAM{]}-CCTTAAACTTTGGAGCCTACCCGCCTG-{[}TAMRA{]})

24\textasciitilde{}27 bp

(Jo et al., 2020)

Biodiversity survey

Hainan Island,China

12S

batra\_F batra\_R batra\_blk

(5'-ACACCGCCCGTCACCCT-3') (5'-GTAYACTTATGTTACTT-3')
(5'-TCACCCTCCTCAAGTATACTTCAAAGGCA-SPC3I-3')

150 bp

(Li et al., 2021)

Biodiversity survey

France

12S

batra\_F batra\_R batra\_blk

(5'-ACACCGCCCGTCACCCT-3') (5'-GTAYACTTATGTTACTT-3')
(5'-TCACCCTCCTCAAGTATACTTCAAAGGCA-SPC3I-3')

20\textasciitilde{}60 bp

(Valentini et al., 2016)

Biodiversity survey

California, United States

12S

12S-V5-F 12S-V5-R

(5'-ACTGGGATTAGATACCCC-3') (5'-TAGAACAGGCTCCTCTAG-3')

106 bp

(W. E. Moss et al., 2022)

Biodiversity survey

Brazilian Atlantic Forest

12S

batra\_F batra\_R batra\_blk

(5'-ACACCGCCCGTCACCCT-3') (5'-GTAYACTTATGTTACTT-3')
(5'-TCACCCTCCTCAAGTATACTTCAAAGGCA-SPC3I-3')

52 bp

(Sasso et al., 2017)

\section*{Figures}

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Figure 1. Flow chart of eDNA detection of amphibians

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