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Croaking Science: The toad fly Lucilia bufonivora in common toads
Croaking Science: The toad fly Lucilia bufonivora in common toads
Blow-flies are dipteran flies that evolved 105 million years ago and there are now over 1,000 species occurring in 150 genera in a range of countries worldwide (Figure 1) (McDonagh, 2009). Within the family Calliphoridae there are 80 species which are known to cause myiasis, which is the infestation of live human and vertebrate animals, where the fly larvae feed on the host’s dead or living tissue, liquid body substances, or ingested food (Zumpt, 1967). There are three main types of blow-fly that cause myiasis. The first are known as saprophages and simply infect animal carcasses that are already dead and decaying. These species do not initiate myiasis but rather take advantage of animals that are already decaying. The second group of blow-flies are generally ectoparasites (i.e. parasites that attach to the skin or fur of an animal) that occasionally initiate myiasis by feeding on damaged tissue of their host. These flies sometimes feed on dead decaying matter, like the saprophages. The last group of blow-flies are known as obligate parasites, only feeding on the living tissue of their host and initiating myiasis as a result (Stevens & Wall, 1997). It has been proposed that the obligate myiasis-causing blow-fly parasites have evolved from an ancestral saprophageous stage, with flies occasionally being attracted to dead or decaying tissue or wounded animals. This later involved into flies which relied more on living tissue until obligate parasites evolved (Zumpt, 1967; McDonagh, 2009).
The blow-fly genus Lucilia comprises approximately 27 species, all of which are similar in appearance. The larvae of the majority of these species are saprophageous, feeding on dead matter. However, there are several species which are occasionally ectoparasites on large mammal species and which may cause myiasis, particularly in sheep (Stevens & Wall, 1997). One species, Lucilia silvarum, sometimes feeds on dead amphibian species, but the toad fly, Lucilia bufonivora, is a highly specialised blow-fly and is an obligate parasite of live prey. Adult L. bufonivora make their first emergence in May to July and seek out living common toads (Bufo bufo) and their lays eggs on the back and flanks of an amphibian host. The larvae then hatch and migrate to the head and nasal cavities of the toad where they continue to feed on the live host (Figure 2). Infected toads tend to change their behaviour and instead of hiding away in sheltered areas, move out into exposed locations where they are susceptible to further infestation by blow-flies. The nasal cavities and head are gradually consumed until eventually the infected toad dies. Once the toad has died, the larvae continue to feed on the flesh of the toad before dropping off to pupate in the soil. Here they undergo metamorphosis and hatch into new adult flies a few weeks later. Toad flies are capable of having three generations each year, depending on the weather and availability of toads.
Toad-flies have a wide distribution across Europe, North Africa and Asia and are particularly common in the Netherlands where between 15 and 70% of common toads may be affected each year, with adults being most commonly affected (Weddeling & Kordges, 2008). Despite this high level of infection, there is no evidence of toad flies causing decline in the common toad. In the UK, toad flies are relatively uncommon and the number of reports each year is low.
L. bufonivora and another blow-fly species L. silvarum are highly similar in appearance. In Europe, L. silvarum tends to only feed on carrion with a preference for dead toads. However, in North America, the species has been recorded as infecting living toads with larvae found in the neck, legs and parotid glands (Eaton et al., 2008). Research by the University of Bristol and Exeter, in collaboration with RAVON, has looked at how closely related L. bufonivora and L. silvarum are to each other. The two species are so similar in appearance it is possible that the eggs found on toads in Europe are from one or both species. Using genetic analysis, the researchers found that the two are sister species, being genetically distinct, but are very closely related and have only recently diverged as separate species. In addition, the researchers found that L. silvarum blow-flies in North America are more closely related to toad flies L. bufonivora, than they are to their own L. silvarum species in Europe. This suggests that obligate parasitism in Lucilia blow-flies may have evolved independently several times and originally diverged from L. silvarum (Arias-Robledo et al., 2008). The obligate parasite traits of L. bufonivora may have evolved as the two species diverged. The findings from this research also show that in Europe and the UK common toads are only infected by L. bufonivora and L. silvarum has yet to become an obligate parasite in these countries (Arias-Robledo et al., 2008). Further research is required to determine the evolutionary status of other closely related blow-fly species such as L. elongata, which is relatively poorly understood.
References
Arias-Robledo, G., Stark, T., Wall, R.L. & Steven, J. R. (2018) The toad fly Lucilia bufonivora: its evolutionary status and molecular identification. Medical and Veterinary Entomology, doi: 10.1111/mve.12328.
Eaton, B. R., Moenting, A. R., Paszkowski, C. A. & Shpeley, D. (2008) Myiasis by Lucilia silvarum (Calliphoridae) in Amphibian Species in Boreal Alberta, Canada. Journal of Parasitology, 94 (4): 949 – 952.
McDonagh, L. M. (2009) Assessing patterns of genetic and antigenic diversity in Calliphoridae (blowflies). PhD thesis, University of Exeter.
Stevens, J. & Wall, R. (1997) The evolution of ectoparasitism in the genus Lucilia (Diptera: Calliphoridae). International Journal of Parasitology, 27 (1): 51-59.
Wellling, K. & Kordges, T. (2008) Lucilia bufonivora-Befall (Myiasis) bei Amphibien in
Nordrhein-Westfalen – Verbreitung, Wirtsarten, Ökologie und Phänologie. Zeitschrift für Feldherpetologie, 15: 183–202.
Zumpt F. & Ledger J. (1967) A malign case of mylasts caused by Hemipyrellia fernandica (Macquart) (Diptera Calliphoridae) in a cape hedgehog (Erinaceus frontalis A. Smith). Acta Zoologica et Pathologica Antverpiensia, 43: 85-91.
Croaking Science: Overwintering in Frog Tadpoles
Overwintering in frog tadpoles
During June and July in the UK the majority of Common Frog (Rana temporaria) tadpoles metamorphose into juveniles which leave ponds for terrestrial habitats. They will spend the rest of the summer and autumn foraging and feeding on small invertebrates in preparation for the winter. However, a number of common frog tadpoles each year will remain in ponds and spend the winter in the water, metamorphosing into juveniles the following spring (Figure 1). What causes some tadpoles to metamorphose and others to remain in the water over the winter? Often, ponds drying out or freezing over can limit the capacity for frog tadpoles to overwinter. However, there are many ponds in the UK where this does not occur, which allows for overwintering of tadpoles. Our understanding of the exact factors which trigger overwintering in tadpoles are not fully understood, but research by Walsh et al. (2016) found that temperature and food availability were key factors. In addition, the decision on whether to over‐winter as a tadpole appears to be made relatively early in the season (Walsh et al., 2008). Under laboratory conditions, lower temperatures and reduced food availability during the summer resulted in a higher proportion of individuals remaining as tadpoles during the winter (Walsh et al., 2016). These findings suggest that cold weather conditions in the early autumn may affect tadpole development, perhaps by affecting endocrine function. However, temperature alone does not appear to be enough to trigger overwintering in tadpoles; food availability also appears to be important. Although these two factors play an important role in triggering overwintering in common frog tadpoles, there are other factors, not fully understood, that appear to be involved.
Living in areas with variations in altitude provides challenges for the development of common frog tadpoles. Higher altitude ponds experience lower temperatures and food availability and these may promote a higher incidence of overwintering in tadpoles. However, Muir at al. (2014) found that in tadpoles living at high altitude in Scotland, individuals had a lower resting metabolic rate which allowed for more energy to be allocated to growth. This is likely to allow individuals to grow faster under cooler environmental conditions and allow tadpoles to metamorphose earlier than expected and prevent the necessity to overwinter. However, if tadpoles developing at high altitudes do need to overwinter, they are able to better tolerate the colder winter temperatures. Muir et al. (2014) found that the tadpoles of Common Frogs at high altitude were able to tolerate freezing for short periods of time. This adaptation provides common frog tadpoles with the ability to withstand the cooler pond conditions at high altitude over the winter and therefore lead to greater survival.
Across Central Europe the incidence of overwintering tadpoles varies by species and is more typical in later breeding species with large tadpoles such as the Midwife Toad Alytes obstetricans, Spadefoot Toad Pelobates fuscus, and water frogs (Pelophylax species) (Gilbert & Harmsel, 2016). The incidence of overwintering in Common Frogs Rana temporaria is rare and was reported for the first time in the Netherlands in 2013 (Gilbert, 2016). This was likely to be due to the unusual weather conditions of 2013/14 along with the sheltered site where this was observed. Overwintering in the Edible Frog (Pelophylax esculentus complex) is a relatively rare phenomenon across Central Europe (Figure 2). Research by Péntek et al. (2018) found that overwintering by tadpoles only seems to occur in unusual weather conditions. In one particular incidence cold temperatures over the winter resulted in late breeding by adults which led to tadpoles getting a late start to their growth and development. Cool conditions early in the autumn (October) appeared to trigger halting of development and the subsequent mild winter promoted successful overwintering by tadpoles of this species (Péntek et al., 2018). Although still relatively rare, overwintering may become more common across Central Europe with changing environmental conditions causing increased variability in seasonal temperatures.
In the United States, 15 species of Ranid frogs occur but only five of these species are known to overwinter as larvae (North American Bullfrog Rana catesbeiana, Pig Frog R. grylio, River Frog R. heckscheri, Red-legged Frog R. aurora, and Southern Mountain Yellow-legged Frog R. muscosa). It appears that pond desiccation is one of the main factors limiting Ranid frog species in North America from overwintering as tadpoles. The Southern Leopard Frog Rana sphenocephala is a common frog inhabiting freshwater ponds across the Northern United States (Figure 3). Tadpoles of this species will remain in the larval stage until reaching a critical metamorphic size and if this is not reached by the end of the autumn, then the tadpoles will overwinter (Pintar & Resetarits, 2018). If winter conditions are suitable, tadpoles will remain in the pond for several years until the critical size for metamorphosis has been reached. This is different to Common Frogs in the UK which do not need a critical size to metamorphose but do require specific environmental conditions. These results highlight the variations that occur across Ranid species in their requirements for metamorphosis.
References
Gilbert, M.J. & Harmsel, R. (2016) Hibernating larvae of the common frog (Rana temporaria) in the Netherlands. Herpetology Notes, 9: 27-30.
Muir, A.P., Biek, R., & Mabel, B.K. (2014) Behavioural and physiological adaptations to low-temperature environments in the common frog, Rana temporaria. BMC Evolutionary Biology, 14: 110.
Péntek, A.L., Sárospataki, M., & Zsuga, K. (2018) Larval overwintering of the Pelophylax esculentus complex in a sodic bomb crater pond near Apaj, Hungary. North-western Journal of Zoology, 2018: e1775023/9.
Pintar, M.R. & Resetarits Jr., W.J. (2018) Variation in pond hydroperiod affects larval growth in Southern Leopard Frogs, Lithobates sphenocephalus. Copeia, 106 (1): 70–76.
Walsh, P.T., Downie, J.R. & Monaghan, P. (2016) Factors affecting the overwintering of tadpoles in a temperate amphibian. Journal of Zoology, 298 (3): 183-190.
Walsh, P.T., Downie, J.R. & Monaghan, P. (2008) Larval over‐wintering: plasticity in the timing of life‐history events in the common frog. Journal of Zoology, 276 (4): 394-401.
Work Experience Blog – Georgia
My name is Georgia, and although my placement with Froglife was not an option I had picked, their charity surrounded British wildlife and reflected my choices to do with animals. I didn’t expect to be completely happy with what I got. However, I was surprisingly interested in what they offered. The people I worked with were friendly and helpful when I asked questions, and my employer aimed to have me outdoors as much as possible so I wasn’t cooped up in the office. I’m usually the kind of person to stay indoors no matter how nice the weather is, but the days I spent working outside were some of the best!
On my first day I was in the office, using a computer to transfer information from forms into a database. I made my way through about half of them and left the rest to finish on another day. After my lunch break, I sorted through children’s drawings of reptiles, amphibians and wildlife to scan into the computer and then spent the rest of the day editing conference videos to be uploaded to YouTube at a later time.
I had to arrive early to work on Tuesday because I was travelling to a school in Whittlesey with Cat and Michelle. They gave talks to Year Fives and Sixes and I walked around the room helping them make their model wildlife gardens. On Wednesday I could have gone with Cat and Michelle to help them with ‘Swimming with Dragons’ pool games they were organising, but I wouldn’t have fit in the car with all of the pool floats, so I stayed in the office and finished editing the conference videos.
On Thursday, I travelled to Boardwalks Nature Reserve with Stuart. For the first half of the day he showed me around, pointing out the different ponds and he even caught a baby smooth newt (an eft) to show me. It was a lot smaller than I originally thought newt efts were – only the size of one section of your finger. After lunch we picked up someone else to help us cut some of the grass. Most of it was extremely overgrown but we managed to cut some paths and we raked the grass into piles for wildlife to live in.
On Friday, I travelled with Michelle to a Nature Reserve to be shown around. We were supposed to be finding a pond in the woods there but on the way we were checking under mats that had been left as places for reptiles to sunbathe. We didn’t find the pond, but on the way back we saw an adder curled up in some grass.
The next week on Monday and Tuesday I went to Hampton with Cat and Michelle to do activities with Year Seven students. The children learnt about amphibians and reptiles and then were separated into three groups and each group took turns trying bug hunting, pond dipping and tree measuring. Although it was cooler under the trees, the heatwave wasn’t pleasant and the children were eager to get back in to school.
Tuesday’s Year Sevens were a lot more interested in Froglife. They found twice as many species of insects as the first group, winning the competition between them. The heat was more bearable and there were less annoying insects like mosquitoes, which may have contributed to a better day.
On Wednesday, I travelled with Stuart to an allotment and a community garden. In the allotment, we gently raked some of the algae on the surface of the pond to the edge to leave some clear areas where sunlight and oxygen could get through. Then we crouched near the disturbed algae and picked out the newt efts that had been raked up with it. Altogether, we found 30 smooth newts and 15 great crested newts, which were a lot larger than the smooth newts. At the community garden, we picked up dead algae around a pond and then we drove back to Froglife for lunch. I scanned some drawings of reptiles I had done into a computer and wrote social media captions for them, then wrote more captions including facts for pictures of wildlife already on the computer.
On Thursday I spent the first half of the day writing my blog post as it was my second-to-last day. Later I was outside unscrewing some screws from a bench that had to be fixed. Then I made a lizard from coloured pipe cleaners to go with a frog and a great crested newt that had already been made. After I finished it I took some pictures – here’s my lizard:
For the first half of Friday I travelled to Boardwalks Nature Reserve with Stuart again to put down more mats for reptiles to sunbathe on. We dug holes in the ground to install wooden posts that the mats were attached to, but the ground was very hard and dry from the heat and lack of rain. This made it very hard to dig, so we only got 4 posts installed before going back for lunch. Then I finished writing the rest of my blog post!
I know I will miss my time at Froglife and the people I’ve met there. It was definitely a positive work experience and although I know a lot of work environments are different from this one, I enjoyed the insight into what it can be like.
Inspired By Nature: A Poem of Return
Inspired by Nature: A Poem of Return
Froglife’s Communications & Fundraising Officer, Ashlea Mawby, felt inspired by the work she does here at Froglife and wrote this poem.
Croaking Science: eDNA for detecting great crested newts – a replacement for traditional survey techniques?
eDNA for detecting great crested newts – a replacement for traditional survey techniques?
Environmental DNA, or eDNA, is released by most organisms as they occupy different habitats. Each species has a unique type of eDNA which can be recognised through laboratory analysis. This provides a novel tool for detecting species within the environment. Sources of eDNA may originate from sloughed skin or hair, eggs, faeces and saliva (Figure 1). Within aquatic habitats, most organisms release eDNA into the surrounding water. An increasing number of methods are now available for detecting the eDNA of a range of aquatic organisms including fish, damselfly nymphs, crustaceans and amphibians. In recent years, techniques have advanced to allow the detection of eDNA of great crested newts from their breeding ponds. This has proved particularly useful for ecologists and voluntary surveyors who need to determine the presence or absence of great crested newts for the purposes of conservation and mitigation. The eDNA technique, though expensive, is usually highly reliable and effective at detecting great crested newts. Research carried out by Biggs et al. (2015) of 35 ponds in Hampshire and Wales showed that eDNA could successfully detect the presence of great crested newts in 99.3% of ponds. This was significantly higher than the success rate of more traditional survey techniques.
Traditional survey techniques for detecting great crested newts such as egg-searching, night torching and bottle trapping are labour intensive, carry certain risks (e.g. potential suffocation of newts in traps) and are not always highly effective. Biggs et al. (2015) found that bottle trapping detected great crested newt presence in 76% of ponds and egg searching in only 44% of visits. Using traditional survey techniques, Natural England advice recommends four visits to a breeding pond in the newt breeding season (mid-March to mid-June) using a minimum of three traditional survey techniques (Natural England, 2015) (Figure 2). The eDNA technique, by contrast, can detect great crested newts on just one visit with minimal disturbance to breeding ponds (Biggs et al., 2015). Therefore, should eDNA replace traditional surveys techniques to detect great crested newts? In this article we highlight the relative advantages of disadvantages of eDNA and point out the limitations that ecologists and voluntary surveyors need to be aware of when using the technique.
One of the major advantages of eDNA is the relative ease that samples can be taken from a pond and the subsequent reduction in labour costs. For example, one fieldworker may be able to take samples for analysis within one survey visit, compared to the multiple sessions required using traditional survey techniques. However, the analysis costs associated with eDNA may be significant, especially if carrying out many samples. A second major advantage of eDNA is its overall reliability compared to other traditional methods. However, a range of studies have found variations in the success of eDNA, ranging from 60% to 99% (Buxton et al., 2017a).
Despite is apparent reliability and success in detecting great crested newts, there are a number of limitations of using eDNA to survey ponds. Six of the main limitations are outlined below:
- Estimation of abundance: the level of eDNA detected from samples taken from a pond does not correlate well with the number of newts in the pond. Biggs et al. (2015) found that at low levels of detection, eDNA may reflect the number of newts but at higher levels, there was only a weak, non-significant correlation. Therefore, it is not currently possible to use eDNA to estimate the population size or relative abundance of great crested newts using ponds. Traditional survey techniques, such as night torching or bottle trapping, may provide an estimate of relative abundance. For example, if 120 newts are captured in bottle traps, this shows that at least this number of newts were present in the pond on this survey occasion.
- Life stages: eDNA only records presence, or recent occupancy, of ponds by newts; it does not show which life stages are present e.g. adults, eggs or larvae, or when they were last in the pond (Rees et al., 2014a). Traditional survey techniques can identify actual animals which is useful for looking at life stage, sex ratio and potentially the body condition of individuals. Also, traditional techniques can record other species of newt which are present in ponds e.g. smooth or palmate newts, which is often important for survey records and provides an indication of the species diversity within a pond.
- Breeding or non-breeding? It is often useful to determine whether great crested newts are breeding within a pond as this may show persistence of a viable, long-term population. However, the eDNA techniques is not able to detect if newts are laying eggs within a pond; it can only determine if they are present, or have recently been present. The traditional survey technique of egg-searching is able to detect the presence of breeding females demonstrating the value in using this technique.
- False absences: a false absence is when a particular technique fails to determine the presence of the species, even though it is present. Although the level of false absences using eDNA was very low in one study carried out by Biggs et al. (2015), this was only carried out on 35 ponds in two counties in the UK. Similar research carried out by Rees et al. (2014) on 38 ponds showed that eDNA had a lower success rate of 89% in detecting great crested newts in ponds. Given the number of false absences in this study Rees et al. (2014) conclude: “Species detection by eDNA and field work is likely to be imperfect and may lead to an underestimation of the distribution of a species especially in the case of rare or threatened species”. False absences may come from a number of sources and are more likely to be a problem when they operate together (Biggs et al., 2015). Common sources of false absences include:
- when there are very few newts in a pond and the water samples fail to collect any eDNA;
- in ponds where surveyors can only access a small proportion of the pond perimeter due to logistical issues;
- at sites which have a very large area of shallow water, which prevents surveyors from reaching the deeper edges often favoured by newts (Biggs et al., 2015) (Figure 3).
- False positives: detecting newts when they are not present is less commonly a problem with eDNA for great crested newts but can occur when there is contamination within the samples collected (Thomsen & Willerslev, 2015). Potential sources of contamination include: excrement from animals that prey on great crested newts; the eDNA of dead great crested newts being carried to the pond by other species (e.g. birds); or unsterilized equipment contaminating samples (Bohmann et al., 2014).
- Reduction or loss of DNA in the water: There are three potential processes that can result in the loss or reduction in the quantity of DNA from within samples: inhibition, degradation, and binding to sediments. This can lead to increases in the likelihood of false absences occurring as less eDNA may be available to detect in analysis.
- Inhibition: eDNA inhibition occurs when factors in the environment, such as decaying organic matter, interfere with the laboratory analysis, resulting in a failure to detect the eDNA (Buxton et al., 2017a).
- Degradation: once eDNA is released into the environment it starts to decay and this rate depends on a range of factors such as UV radiation, temperature and microbes which occur in the pond (Buxton et al., 2017; Rees et al., 2017). Currently, our understanding of how quickly eDNA degrades under different conditions is limited (Rees et al., 2014a). This is of importance since if eDNA degrades faster in some ponds than others, this may limit the likely time window when surveys can be carried out. Although research has shown that eDNA may be present in ponds in all months of the year (Rees et al., 2017), the realistic time frame for detecting adults is March to the end of June (Rees et al., 2017) which is the same as for traditional survey techniques. However, if ponds contain breeding adults and subsequent larvae, these may be detected in ponds up until the end of August (Buxton et al., 2017).
- Binding to sediments: research has also demonstrated that eDNA will bind to certain sediments in ponds. Buxton et al. (2017a) have shown that eDNA binds more effectively to clay and organic soils when compared to sand. This has important consequences for surveying since the eDNA will disappear much more quickly in ponds with clay sediments. This means that if great crested newts are present in two ponds, one with a clay sediment and one with a sand sediment, the available time frame for surveying may be shorter in the pond with clay lining. A late survey carried out may result in a false absence record in the pond with clay sediment, even though the newts had recently been present. Given the variation in occurrence of eDNA in water samples, the potential for false absences needs to be fully understood before using eDNA as a survey tool and it is important to realise that eDNA research is still an evolving discipline (Buxton et al., 2017a).
Overall, although using eDNA to detect the presence of great crested newts is highly effective and usually reliable, ecologists and surveyors must be aware of the potential limitations when carrying out surveys. Therefore, caution should be taken when analysing the results from eDNA; ecologists and surveyors should consider eDNA as an additional technique to complement traditional survey methods, rather than being viewed as replacing existing techniques. We conclude this article with a statement by Rees at al. (2014a): “Environmental DNA methodologies should not be used to replace or disregard the knowledge and expertise of experienced field ecologists and taxon specialists, but should become an important tool to enhance limited conservation resources”.
References
Biggs, J., Ewald, N., Valentini, A., Gaboriaud, C., Dejean, T., Griffiths, R.A., Foster, J., Wilkinson, J.W., Arnell, A., Brotherton, P., Williams, P. & Dunn, F. (2015) Using eDNA to develop a national citizen science-based monitoring programme for the great crested newt (Triturus cristatus). Biological Conservation, 183: 19–28.
Buxton, A.S., Groombridge, J.J., Zakaria, N.B. & Griffiths, R.A. (2017) Seasonal variation in environmental DNA in relation to population size and environmental factors. Science Reports, 7: 46294; doi: 10.1038/srep46294.
Buxton, A.S., Groombridge, J.J. & Griffiths, R.A. (2017a) Is the detection of aquatic environmental DNA influenced by substrate type? PLoS ONE, 12 (8): e0183371.
Bohmann, K., Evans, A., Thomas, M., Gilbert, P., Carvalho, R., Creer, S., Knapp, M., Yu, D.W. & de Bruyn, M. (2014) Environmental DNA for wildlife biology and biodiversity monitoring. Trends in Ecology & Evolution, 29 (6): 358-367.
Natural England (2015) Great crested newts: surveys and mitigation for development projects. https://www.gov.uk/guidance/great-crested-newts-surveys-and-mitigation-for-development-projects#when-to-survey. Accessed 20th June 2018.
Rees, H.C., Bishop, K., Middleditch, D.J., Patmore, J.R.M., Maddison, B.C. & Gough, K.C. (2014) The application of eDNA for monitoring of the Great Crested Newt in the UK. Ecology and Evolution, 4 (21): 4023–4032.
Rees, H.C., Maddison, B.C., Middleditch, D.J., Patmore, J.R.M. & Gough, K.C. (2014a) 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.
Rees, H.C., Baker, C.A., Gardner, D.S., Maddison, B.C. & Gough, K. (2017) The detection of great crested newts year round via environmental DNA analysis. BMC Res Notes, 10: 327. DOI 10.1186/s13104-017-2657-y.
Thomsen, P.F. & Willerslev, E. (2015) Environmental DNA – An emerging tool in conservation for monitoring past and present biodiversity. Biological Conservation, 183: 4–18.