Environmental DNA
(eDNA) for Biodiversity Monitoring in Cave Ecosystems
Mike Buchanan 2025
Abstract
Keywords: environmental DNA, caves, biodiversity monitoring,
groundwater, metabarcoding, conservation
1. Introduction
Cave ecosystems host specialised fauna, including
stygobionts, troglobionts, and species of high conservation concern.
Traditional survey methods often under-detect these organisms due to their
cryptic behaviours and inaccessible habitats. eDNA offers a non-invasive
alternative, capable of detecting rare and elusive taxa at high sensitivity
(Saccò et al., 2022). However, successful application in caves requires careful
attention to study design, environmental constraints, and integration with
conventional approaches.
2. Accuracy and Effectiveness of eDNA in Caves
Research indicates that eDNA effectively detects aquatic
cave species such as fish, crustaceans, and amphibians, often outperforming
visual surveys (Boyd et al., 2020). For terrestrial cave fauna, success is
contingent on the sampling substrate: guano, sediments, and biofilms yield more
reliable results than water alone (Saccò et al., 2022). While eDNA provides
high sensitivity, accuracy may be reduced by false positives from contamination
or transported DNA, and false negatives where biomass is very low (Goldberg et
al., 2016).
3. Sampling Effort and Duration
Standard practice recommends three to five replicates of
water (1–4 L filtered per replicate) or sediment samples at each site. Temporal
replication is equally important, with two sampling events considered minimal
and three to four across seasonal periods providing more robust coverage (Jerde
et al., 2019). Field campaigns can be completed within days, although
laboratory processing and bioinformatics can extend timelines to several weeks.
4. Optimal Sample Types for Cave eDNA
Different sample matrices offer varying strengths:
·
Water samples: suited to aquatic organisms (Boyd
et al., 2020).
·
Sediment samples: concentrate DNA, preserving it
longer and increasing detection rates for benthic taxa (Saccò et al., 2022).
·
Guano: effective for bats and cave-roosting
birds, also useful in dietary studies (Saccò et al., 2022).
·
Biofilms and wall scrapes: retain DNA from
surface-associated taxa (Saccò et al., 2022).
Groundwater samples: informative for aquifer-connected fauna
but require caution due to DNA transport (Korbel et al., 2024).
5. Practical Implementation of eDNA Studies in Caves
A typical eDNA workflow in caves includes:
·
Planning: Define targets, select markers, and
design replication (Goldberg et al., 2016).
·
Field collection: Sterile sampling of water,
sediment, or guano with inclusion of field blanks.
·
Preservation: On-site filtration where possible,
or storage in ethanol/preservative.
·
Laboratory analysis: DNA extraction, PCR
amplification with species-specific or metabarcoding primers, sequencing, and
bioinformatics.
·
Data interpretation: Apply rigorous controls,
quality filters, and statistical models such as occupancy models for reliable
inference (Goldberg et al., 2016).
·
Multiple sampling events — typically two to four
— are recommended to account for seasonal or hydrological variation.
6. Environmental Factors Affecting eDNA Accuracy
Factors that reduce detection reliability include:
·
Low organismal abundance and DNA shedding rates
(Saccò et al., 2022).
·
Hydrological transport and dilution in
groundwater (Korbel et al., 2024).
·
PCR inhibitors in cave sediments and guano (Boyd
et al., 2020).
·
DNA degradation from microbial activity or
warmer microclimates (Jo et al., 2020).
·
Limited barcode representation in reference
databases (Saccò et al., 2022).
7. Persistence of eDNA in Cave Environments
eDNA persistence varies by matrix:
·
Water: generally, days to weeks in pools (Jo et
al., 2020).
·
Groundwater: weeks to over a month in stable,
low-microbial conditions (Korbel et al., 2024).
·
Sediment/guano: months to years, though this may
reflect historical presence rather than current occupancy (Saccò et al., 2022).
8. Integration with Traditional Survey Methods
·
eDNA from guano with acoustic surveys enhances
bat monitoring (Saccò et al., 2022).
·
eDNA complements netting and trapping for
aquatic invertebrates and fish (Boyd et al., 2020).
·
Traditional captures provide voucher specimens,
improving local barcode databases for eDNA interpretation (Saccò et al., 2022).
·
This complementary approach maximises detection
accuracy, provides ecological context, and supports species management
decisions.
9. Conclusion
eDNA represents a transformative tool for cave biodiversity
monitoring. When applied with robust sampling design, careful substrate
selection, and complementary traditional methods, it offers a powerful means of
detecting both aquatic and terrestrial cave fauna. Despite challenges linked to
DNA persistence and environmental variability, its integration with
conventional survey methods ensures comprehensive biodiversity assessments in
these critical and understudied ecosystems.
References
Boyd, S.H., Fensome, R.A., Niemiller, M.L. & Hollenbeck,
C.M. (2020) Detection of an endangered cave crayfish using environmental DNA.
Freshwater Biology, 65(9), 1546–1554.
Goldberg, C.S., Strickler, K.M. & Pilliod, D.S. (2016)
Moving environmental DNA methods from concept to practice for monitoring
aquatic macroorganisms. Biological Conservation, 183, 1–3.
Jerde, C.L., Mahon, A.R., Chadderton, W.L. & Lodge, D.M.
(2019) eDNA as a conservation tool: Designing effective sampling strategies.
Conservation Genetics Resources, 11(1), 1–10.
Jo, T., Murakami, H., Yamamoto, S., Masuda, R. &
Minamoto, T. (2020) Effect of water temperature and fish biomass on
environmental DNA shedding, degradation, and size distribution. Ecology and
Evolution, 10(19), 10023–10032.
Korbel, K.L., Hose, G.C., Carini, G. & Seymour, J.R.
(2024) Detection, movement and persistence of invertebrate eDNA in subterranean
systems. Molecular Ecology Resources, 24(2), 211–225.
Saccò, M., Watson, G., Lunghi, E., et al. (2022) eDNA in
subterranean ecosystems: Applications, technical challenges, and prospects.
Environmental DNA, 4(5), 836–853.
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