Environmental DNA (eDNA) for Biodiversity Monitoring in Cave Ecosystems

Mike Buchanan - 2025 - 2 of 3

Abstract

Environmental DNA (eDNA) is an increasingly applied molecular tool that enables detection of organisms through traces of genetic material shed into the environment. Subterranean ecosystems such as caves present unique opportunities and challenges for eDNA monitoring due to their stable microclimates, complex hydrology and presence of cryptic species. This paper reviews the accuracy and effectiveness of eDNA in caves, explores optimal sampling designs, highlights environmental factors affecting detection and discusses integration with traditional ecological methods. Recommendations for best practice are provided, positioning eDNA as a complementary approach to conventional cave biodiversity surveys.

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 subterranean 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 at cost.

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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