Environmental DNA (eDNA)

Environmental DNA (eDNA)

grooming 3A6A5458 (Killeen)Aquatic animals release DNA into the water when they excrete urine or faeces or slough skin cells. The platypus is expected to be a good source of such DNA as it habitually urinates, defaecates and grooms its fur while swimming or floating. In addition, the platypus’s front feet and bill are covered by bare skin that may rub against rocks or submerged branches as an animal forages. In contrast, studies to date indicate that eDNA is not a very effective way to detect Australian water-rats (or rakali), which are much more likely to groom and defaecate on land as compared to a platypus.

Through the magic of modern genetics, water samples can be processed to extract and amplify whatever environmental DNA (or eDNA) is present and then identify its source with considerable accuracy.

a 5.5km upstrm of bridge look u.s.Because life isn’t meant to be easy, eDNA is both consumed by microbes and degraded by the ultraviolet (or UV) component of sunlight. It can also become unavailable for sampling after settling to the bottom of a water body or becoming adsorbed onto particles of sediment or other materials in the water column. In practice, eDNA becomes undetectable in natural water bodies over periods ranging from one day to possibly as much as several weeks (Rees et al. 2014; Goldberg et al. 2016), typically persisting longer in water that is cool, shaded and alkaline as compared to water that is warm, exposed to sun and has a neutral to acid pH (Strickler et al. 2015). These characteristics can in turn limit use of eDNA as a reliable technique to monitor how disturbance events affect platypus activity. For example, water temperature, the rate of microbial decomposition and the amount of light reaching the water have all been found to increase substantially after bushfire due to reduced tree canopy cover (Rodriguez-Lozano et al. 2015). One would therefore predict that platypus eDNA will be detected less often in badly burnt habitats as an outcome of reduced eDNA persistence even if platypus numbers remain entirely unchanged.

The concentration of eDNA in flowing waters is also affected by how rapidly it’s carried downstream, which in turn depends on channel morphology and flow volume and velocity (Pilliod et al. 2013). There seems to be an emerging consensus that eDNA degrades or otherwise becomes undetectable quite rapidly (at least in flowing waters), so its detection can reasonably be inferred to reflect recent activity by the detected species near the sampling site (e.g. Spear et al. 2015; Wilcox et al. 2016). However, eDNA’s known sensitivity to multiple factors (including seasonal variation in a species’ activity or behaviour) means that conclusions derived for one time and place can’t necessarily be assumed to apply to other habitats or seasons (Shaw et al. 2016), with significant day-to-day variation in eDNA concentration sometimes occurring even at a given site (Tillotson et al. 2018).

Based on field work conducted along small streams near Melbourne, Lugg et al. (2017) concluded that a 95% platypus eDNA detection probability can be achieved by analysing two water samples (with two replicates tested per sample) using qPCR methodology. The sensitivity of eDNA as a detection method for platypus was also estimated to be roughly 2-4 times better than when fyke nets were used for the same purpose. However, this finding is at best a starting point for comparing the relative sensitivity of the two methods, given that the frequency of platypus captures in nets can vary by a factor of 4 times or more depending on how the nets are set.

By the same token, more research is needed to develop guidelines to optimise detection of platypus eDNA across the full range of habitats and flow regimes associated with these animals, particularly at sites where they occur in low numbers. Meanwhile, as a useful rule, the best time of year to sample platypus eDNA (especially to minimise the risk of false negative outcomes) is likely to be when animals are most active and mobile, e.g. late winter to mid-spring in Victoria (Serena and Williams 2012).

Photos: APC (below) and courtesy of Ann Killeen (above)

Literature cited

Goldberg, C. S., et al. (2016). Critical considerations for the application of environmental DNA methods to detect aquatic species. Methods in Ecology and Evolution 7, 1299-1307.

Lugg, W. H., Griffiths, J., van Rooyen, A. R., Weeks, A. R., and Tingley, R. (2018). Optimal survey designs for environmental DNA sampling. Methods in Ecology and Evolution 9, 1049-1059.

Pilliod, D. S., Goldberg, C. S., Arkle, R. S., and Waits, L. P. (2013). Estimating occupancy and abundance of stream amphibians using environmental DNA from filtered water samples. Canadian Journal of Fisheries and Aquatic Sciences 70, 1123-1130.

Rees, H., Maddison,B. C., Middleditch, D. J., Patmore, J. R. M., and Gough, K. C. (2014). The detection of aquatic animal species using environmental DNA – a review of eDNA as a survey tool in ecology. Journal of Applied Ecology 51, 1450-1459.

Rodriguez-Lozano, P., Rieradevall, M., Rau, M. A., and Prat, N. (2015). Long-term consequences of a wildfire for leaf-litter breakdown in a Mediterranean stream. Freshwater Science 34, 1482-1493.

Shaw, J. L. A., Clarke, L. J., Wedderburn, S. D., Barnes, T. C., Weyrich, L. S., and Cooper, A. (2016). Comparison of environmental DNA metabarcoding and conventional fish survey methods in a river system. Biological Conservation 197, 131-138.

Spear, S. F., Groves, J. D., Williams, L. A., and Waits, L. P. (2015). Using environmental DNA methods to improve detectability in a hellbender (Cryptobranchus alleganiensis) population. Biological Conservation 183, 38-45.

Strickler, K. M., Fremier, A. K., and Goldberg, C. S. (2015). Quantifying effects of UV-B, temperature and pH on eDNA degradation in aquatic microcosms. Biological Conservation 183, 85-92.

Tillotson, M. D., Kelly, R. P., Duda, J. J., Hoy, M., Kralj, J., and Quinn, T. P. (2018). Concentrations of environmental DNA (eDNA) reflect spawning salmon abundance at fine spatial and temporal scales. Biological Conservation 220, 1-11.

Wilcox, T. M., McKelvey, K. S. Young, M. K., Sepulveda, A. J., Shepard, B. B., Jane, S. F., Whiteley, A. R., Lowe, W. H., and Schwartz, M. K. (2016). Understanding environmental DNA detection probabilities: a case study using a stream-dwelling char Salvelinus fontinalis. Ecological Applications 194, 209-216.