Environmental DNA (eDNA)
Aquatic animals release DNA into the water whenever they excrete urine or faeces, slough skin cells or bleed after being injured. 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 which (particularly in the case of the bill) may rub against rocks or submerged branches when an animal forages.
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.
Because life isn’t meant to be easy, eDNA is both consumed by microbes and degraded by the ultraviolet component of sunlight. It can also become unavailable for sampling after settling to the bottom of a water body or becoming adsorbed onto other materials. In practice, eDNA becomes undetectable in natural water bodies over periods ranging from one day to potentially 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).
The concentration of eDNA in flowing waters is also affected by transport in the current, which in turn depends on channel morphology as well as the velocity and volume of flow (Pilliod et al. 2013). There seems to be an emerging consensus that eDNA degrades or otherwise becomes undetectable quite rapidly (at least in the case of 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, the known sensitivity of eDNA concentration to a wide range of factors (local hydrology, habitat attributes, water temperature, seasonal variation in activity and behaviour, etc.) means that conclusions derived from data obtained at a given time and place can’t be assumed to apply to other seasons (Shaw et al. 2016) or across habitats, 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 platypus eDNA detection probability of 95% can be achieved by analysing two water samples (300-500 millilitres/sample) using qPCR methodology (with two replicates tested per sample). The sensitivity of eDNA as a detection method for platypus was also estimated to be roughly 2-4 times better than use of fyke nets for the same purpose. However, this is at best a starting point for comparison given that platypus capture efficiency in nets can vary by a factor of 4 times or more depending on how they 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 (to minimise the risk of false negative outcomes) is predicted 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)
Counts of burrow entrances
Platypus mainly rest in burrows that have platypus-sized entrances opening near the edge of the water. Unfortunately, a number of factors preclude using counts of burrow entrances as an index of platypus abundance:
- Platypus burrow entrances are characteristically very difficult to detect – they can be located underwater and are often concealed by undercut banks or trees, overhanging shrubby vegetation or long grass (as shown at right), piles of miscellaneous woody debris, or even man-made structures such as concrete drains and boardwalks (Serena 1994; Gardner and Serena 1995; Serena et al. 1998; Thomas et al. 2019). For example, the entrances of only 6 of 57 known platypus burrows could be identified with reasonable certainty in a radio-tracking study conducted near Melbourne (Serena et al. 1998).
- Apart from the mothers of small juveniles, a platypus normally will occupy several different burrows scattered across its home range in a period of just a few weeks (e.g. Serena 1994; Gardner and Serena 1995; Gust and Handasyde 1995; Thomas et al. 2019).
- Burrows are occupied sequentially by different individuals over time (Serena 1994; Serena et al. 1998), and presumably may continue to exist for many years (even if a population becomes locally extinct) unless substantial bank erosion or compaction occurs.
- There are currently no known grounds for clearly distinguishing a platypus burrow from a water-rat/rakali burrow – the two species are fairly similar in size, and in fact have been documented to occupy the same burrow sequentially (Serena 1994).
Conventional and sonar-based cameras
The use of land-based camera traps to identify platypus activity is limited by the fact that these animals spend little time out of the water and have a very low profile when swimming on the surface (though see photo at right of a platypus on a log, taken by a camera meant to survey turtles).
Underwater infrared camera systems have been developed to assist with nocturnal fish research but have a limited range of detection even in clear water (e.g. Chidami et al. 2007). Alternatively, it’s now possible to purchase video equipment that uses sound waves instead of light waves to create underwater images. This technology copes well with darkness and high turbidity and registers images over a far greater distance than is possible with conventional video gear. The downside is that it’s quite expensive to install, especially if numerous sites are to be monitored simultaneously.
Photo courtesy of Jesse Miller
Chidami, S., Guenard, G., and Amyot, M. (2007) Underwater infrared video system for behavioural studies in lakes. Limnology and Oceanography: Methods 5, 371-378.
Gardner, J. L., and Serena, M. (1995). Spatial organisation and movement patterns of adult male platypus, Ornithorhynchus anatinus (Monotremata: Ornithorhynchidae). Australian Journal of Zoology 43, 91-103.
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.
Gust, N. and Handasyde, K. (1995). Seasonal variation in the ranging behaviour of the platypus (Ornithorhynchus anatinus) on the Goulburn River, Victoria. Australian Journal of Zoology 43, 193-208.
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.
Serena, M. (1994). Use of time and space by platypus (Ornithorhynchus anatinus: Monotremata) along a Victorian stream. Journal of Zoology 232, 117-131.
Serena, M., and Williams, G. A. (2012). Effect of sex and age on temporal variation in the frequency and direction of platypus (Ornithorhynchus anatinus) captures in fyke nets. Australian Mammalogy 34, 75-82.
Serena, M., Thomas, J. L., Williams, G. A., and Officer, R. C. E. (1998). Use of stream and river habitats by the platypus, Ornithorhynchus anatinus, in an urban fringe environment. Australian Journal of Zoology 46, 267-282.
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.
Thomas, J. L., Parrott, M. L., Handasyde, K. A., and Temple-Smith, P. (2019). Burrow use by juvenile platypuses (Ornithorhynchus anatinus) in their natal home range. Journal of Mammalogy 100, 1182-1190.
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.