The potential to reduce the suffering of animals living in the wild by using eDNA sampling

by Animal_Ethics21 min read16th Nov 2020No comments

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Wild Animal Welfare
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Cross-posted from the Animal Ethics blog.

Introduction

Animals living in the wild suffer for a number of reasons including hunger, predation, and disease. There is a lack of consideration and care for animals living in the wild, despite their ability to suffer the same as domesticated animals and human beings. Environmental DNA (eDNA) sampling is a noninvasive research technique that can be used to learn more about animals living in the wild, such as whether they are suffering from a particular disease. This information could be used in the future to help animals when they are in need. 

eDNA is defined as the DNA that an animal leaves behind in their environment.¹ Common sources of eDNA include hair,2 feathers,3 and feces,4 while less common sources of eDNA include snail trails5 and scent marks.6 

After collection, eDNA samples are replicated by applying the polymerase chain reaction method. This technique duplicates a particular sample of DNA, generates thousands to millions of copies of that DNA, and allows scientists to obtain longer chains that help to reveal new information about animals. For example, species have been identified,7 species population sizes have been estimated,8 diets have been analyzed,9 and diseases have been detected.10 Until recently, invasive techniques have been used for these purposes. However, eDNA sampling offers the same insights into the lives of animals without having to unnecessarily interact with them. 

The information obtained about animals living in the wild by using eDNA sampling could potentially help many animals. In accordance with welfare biology, a proposed research area that would study sentient beings and their environment with respect to the animals’ wellbeing, this article will specifically focus on highlighting the advantages of eDNA and the possible ways it could be used to help alleviate the suffering of animals living in the wild in the near future.

To begin, the article will highlight the dangers associated with live trapping nonhuman animals and how to avoid these dangers by using eDNA sampling, also noting the important advantages that eDNA sampling has over many invasive trapping methods. Next, the article will talk about using eDNA sampling to detect disease in nonhuman animals. The article will also cover using eDNA sampling for species detection. Lastly, the article will talk about some possible ways in which eDNA sampling could be used in the future to help animals living in the wild.

Using eDNA sampling in place of invasive research techniques

Live trapping

Animals living in the wild are often at risk of being trapped and killed by human beings. Often they are killed for food, fur, or to control the number of animals within a geographical area. In addition to these reasons, animals are often trapped in live traps for scientific research. 

Live traps are baited to lure an animal into a situation from which they cannot escape. The traps are not designed to kill the target species. Typical forms of live trapping are leg nooses, mist nets, funnel traps, and boxes used to trap small mammals. 

Live trapping is not usually done to help individual animals, for example, to trap injured animals and help them recuperate or to stop animals from crossing a busy motorway by trapping them to transport them safely across. Instead trapping is usually done to further scientific knowledge about animals. In addition, the animal’s wellbeing is not the highest priority when they are trapped unless it is important to the research taking place. Despite being designed not to kill animals, these traps can cause stress, physical harm, and death in some circumstances. 

Stress from trapping and handling

There have been many documented instances in which trapping and handling have been shown to cause stress in animals. A number of examples are given below. 

Bighorn sheeps have been found to experience increased cardiac activity, indicative of a stress response, for almost two hours after being trapped and handled.11 Stress responses have been exhibited by American red squirrels trapped in live traps, with their stress response worsened when their trap had increased visibility to the outside world. After two hours, their stress hormone concentrations increased further.12 Corticosterone levels in house sparrows increase when they are trapped, and become further elevated after about 15 minutes.13 Voles have also been found to experience capture-related stress.14 Animals in laboratories show elevated stress levels from being handled. A review of 80 studies showed that, on average, the animals experienced elevated heart rate and blood pressure for at least 30 minutes after being handled.15 Lastly, young snails have suffered high mortality rates from the stress of being handled.16

These results show that animals experience a stress response, indicated by elevations in heart rate, corticosterone levels, and blood pressure. This will happen whether or not their wellbeing is taken into consideration by, for example, making the traps less uncomfortable or checking the traps more frequently so that the animals are held for shorter periods of time. Being trapped and handled is a stressful experience for any nonhuman animal, as it would be for a human being.

The psychological effects of trapping and handling can be extreme, and in some cases an animal’s anxiety can lead them to make desperate attempts to escape that result in physical harm, despite the security of the trap. In addition to this, animals sometimes experience physical harm as a consequence of being handled or from a defect in the trap.

Physical harm from trapping

Live trapping and handling are methods designed not to cause physical harm to animals; however many animals are injured while being trapped or handled. A number of documented instances are listed below. 

An analysis of 16 studies was conducted and showed that shrews died as a result of an accident whilst being trapped, on average, 47% of the time.17 Winter hares suffered a significant decrease in their body mass after being trapped and handled for the first time.18 Scientists found  a possible positive correlation with handling albatross chicks raised in captivity and elevated muscle enzyme levels.19 Elevated muscle enzyme levels are indicative of muscle damage and can negatively affect the birds. Trapping and handling caused 15% of captured little bustards to suffer mobility issues and 43% of those birds later died.20

In addition to the physiological and psychological risk-factors associated with handling and trapping animals, it is believed that invasive trapping methods are likely to give inaccurate results regarding the behavior, health, or activity of animals living in the wild, due to the stress they experience. As a result, there have been calls for the reform of animal care policies in research journals and stricter institutional supervision.21 It should be noted that new information about the lives of animals should not be at the expense of an individual animal’s wellbeing.  Research should be conducted noninvasively and techniques such as eDNA sampling should be used whenever possible.

Advantages of using eDNA sampling instead of live trapping

The most obvious advantage of using eDNA sampling is that it is noninvasive and safe for animals. As previously demonstrated, trapping animals for scientific research is invasive and can cause harm to animals, both psychologically and physically. Advocating for the use of eDNA sampling and other effective noninvasive techniques to be used in scientific research is a way of helping animals living in the wild.

Other advantages of eDNA sampling include accuracy of results, cost-effectiveness, and low sampling effort. eDNA sampling may also be used in a variety of different environments. Accuracy, cost-effectiveness, and low sampling effort play an important role in making the technique desirable to the scientific community. 

Accuracy 

eDNA samples are subject to several conditions that will affect the potential for providing information about animals, including the accuracy of the findings.

It is important to assess the quality of eDNA samples as contamination and age of the sample can lead to misleading results. Particularly, the type, temperature, and time since the DNA left an animal will determine the quality of the sample. For example, fecal samples taken in a warm, dry climate that are up to a week old should provide optimal results.22 Additionally, within different environments the optimal periods of time for eDNA sampling will differ greatly from one another. For example, there have been experiments showing that optimal sampling time for eDNA is up to 10 hours in seawater,23 up to 14 days in freshwater,24 and up to 93 days in sediment.25 Another example showed that eDNA samples taken from soil were found to accurately determine the presence of species that were detected by camera imaging between 30 and 150 days beforehand.26

In addition, the success of eDNA sampling is dependent on how much is known about a string of DNA. If information is not available about the DNA sequence of a particular species, then it will be almost impossible to identify key features.

Lastly eDNA has been known to give false positives, reporting a positive result for the presence of an animal who wasn’t there, and false negatives, reporting a negative result when the animal in fact was present. This can be for a number of reasons, but will most likely occur when the sample is contaminated,27 when the sample hasn’t been handled appropriately,28 or when the sample area doesn’t have a high enough concentration of DNA.29 As more is learned about the DNA of individual species and catalogues of known DNA are shared among the scientific community, the accuracy of eDNA sampling will improve.

Despite the concerns regarding eDNA sampling, continuous improvements in the equipment and methodology have revealed an impressive accuracy when analyzing samples. A number of studies have shown eDNA sampling to equal or out-perform other techniques used to gather the same information about animals. Below are two examples.

eDNA samples were taken from two rivers in southern Australia and scientists found that using five 1 liter samples of water for eDNA sampling was sufficient to detect all of the species of fishes that had been trapped using invasive methods.30 When detecting the brown marmorated stink bug, scientists noted that the insects would often eat the peaches, leaving behind ample eDNA on the peach after they had finished. The field scientists found that sampling eDNA from peaches was a considerably more sensitive technique for detecting the stink bug and offered an effective alternative to the two invasive methods previously used.31

While invasive methods like trapping can be improved by decreasing the discomfort of trapped animals, the stress animals will experience is inevitable and will affect the accuracy of the findings. eDNA sampling is not limited in this way and will continue to improve as more DNA is catalogued. Similarly, eDNA sampling will become more cost-effective as it becomes more accessible and widely used.

Cost-effectiveness

Several experiments have shown eDNA sampling to be more cost-effective than an invasive method used to learn the same information. In one example, an experiment compared the cost of eDNA sampling to an invasive method designed to trap fishes. Scientists participating in the experiment calculated that the overall cost of collecting and analyzing eDNA samples came to $678, whereas using some invasive methods cost far more.32

Scientists also compared the cost-effectiveness of eDNA sampling to typical surveying techniques such as torchlight surveys and egg counting. It was found that surveying could cost up to several thousands of dollars whereas eDNA sampling would only cost a few hundred.33

Environment

The results using traditional techniques used to gather information about animals may be affected by the sampling environment. For example, detecting an animal in snow can be difficult because the animal may be camouflaged by their surroundings. eDNA sampling has an advantage over other research techniques in that it can be applied to all environments where DNA exists, and does not require the presence of an animal at the time that the sample is taken.

One of the most popular environments to apply eDNA sampling is water, including oceans and areas of freshwater. The level of success achieved in these environments depends on many features including DNA degradation, species abundance, and the size of the body of water. The majority of published studies about eDNA sampling have concentrated on making discoveries about animals in water, particularly focusing on species detection. An example of one is a review that collated a number of studies focused on detecting aquatic species. They concluded that, compared to other techniques, eDNA sampling allows for more cost-effective, targeted estimates of the number of different species in water.34

Another research team also found that eDNA sampling was very effective in tropical environments. eDNA samples were used to identify different species of frogs in a dry forest region in Bolivia. The research team determined that eDNA sampling was a successful way of detecting different species of frogs in a tropical environment, especially regions with a high density of amphibians, making it more likely that the eDNA sample would contain relevant information.35

Scientists wanting to understand whether eDNA sampling would help them detect species in harsh conditions like snow looked for evidence of the presence of three species in areas of Montana in the US. eDNA samples were taken from a variety of places including snow-tracks, from snow in an area where the animals had previously been captured on camera, and from hair samples collected 1.6 months beforehand. The snow samples were then filtered and the remaining DNA was processed. All three species were successfully detected and eDNA sampling was determined to be far superior to general surveying techniques in a snowy environment.36

Lastly, a review was conducted to determine how well eDNA sampling performed in the Arctic tundra. The molecular-based technique was found to be very effective in this environment, providing information about the diet of the animals living nearby as well as the identification of species and population structures.37

Sampling effort

Sampling effort is a measure of the time and effort that it takes for researchers or technicians to extract DNA samples. eDNA sampling is considered to require less sampling effort compared to other techniques because of its simplicity. Most invasive techniques used to collect data from animals are time-consuming, require many workers, and normally a high level of skill to reduce the harms suffered by animals during trapping. Also, in most cases eDNA sampling requires less equipment and sample-taking is considered to be easier than other research techniques. Below are some examples.

Field technicians found that the collection of water samples for eDNA sampling was simple compared to catching frogs for invasive testing. In addition, the water samples could be collected by most field technicians, whereas taking tissue samples required workers with a higher technical ability and sometimes special certification.38 Taking eDNA samples was also compared against an invasive method used to manipulate the movement of fishes. To apply the invasive technique, the workers had to enter the creek which was far more effort than taking an eDNA sample from the creek bank. In terms of time, eDNA sampling took 6.8 person-hours, whereas the invasive method took both 30 person-hours, and 90 person-hours, depending on the technology of the technique.39 Another study compared the results from eDNA samples taken from a particular area with the results from a 9-year camera trapping study in California. eDNA sampling was able to detect several small species missed by the camera traps and proved to be much easier than camera trapping.40

Overall eDNA sampling has proven to be a simple method to implement, especially when compared to other methods used for purposes such as detecting disease.

Other forms of noninvasive research that can be used instead of live trapping

The use of noninvasive techniques in research are rapidly increasing. Examples of noninvasive techniques, other than eDNA sampling, include thermography, the use of drones, and camera trapping. 

Each technique offers a unique way of understanding animals living in the wild. Thermography is a technique using an infrared camera to observe animals. The camera is normally used in an environment that is colder than the animal’s body temperature, allowing the creation of an outline of the animal’s body shape. Camera-carrying drones have been used for almost a decade to monitor animals in the wild. Camera trapping is a method used to capture videos of animals living in the wild. The camera is usually equipped with a sensor triggered by movement or heat and will begin recording as animals approach it.

For these techniques to be successful, animals are required to be in the presence of someone or something at the time the sample is being taken. In the strictest sense of noninvasive research, if animals are made aware of the alien presence, because of noise or movement from a drone, for example, and change their behavior by running away or investigating the object, it is questionable whether these techniques are really noninvasive. If care is taken not to disturb nearby animals, then this problem does not exist when taking eDNA samples.

Detecting disease using eDNA sampling

Disease is one of the many causes of suffering for animals living in the wild, and it can be difficult to detect. Aside from eDNA sampling, there are no noninvasive ways of definitively detecting disease for these animals.41 Presently disease is detected in a small number of superficial ways. For example, if a group of animals is under observation, and some animals seem to be acting abnormally, if there are visible abnormalities in an animal’s appearance, or if there is a large number of corpses, then this could indicate the presence of disease. 

eDNA sampling provides an option for monitoring the health of animal populations and potentially detecting diseases that involve an observable change in DNA in its early stages, such as viruses or bacterial diseases. In some cases, this could lead to the development of vaccines that can prevent suffering and death among animals. The following are some examples.

There have been recent advancements in the early detection of Squirrel Pox Virus using eDNA sampling in the form of shed squirrel hair samples.42 eDNA sampling in the form of urine and feces, has been proven to be a successful, noninvasive method to identify the transmission of tuberculosis from badgers to cows in the UK.43 Noninvasive urine sampling has been proven to be successful in providing a method of early detection of Brucellosis in dogs.44 eDNA sampling was successful in detecting Ranavirus and it was concluded that eDNA sampling worked better than common invasive testing methods in detecting disease in amphibians.45

Detecting parasites using eDNA sampling

As well as detecting diseases caused by bacteria or viral pathogens, eDNA is also capable of detecting disease caused by parasites. Parasitic diseases can be incredibly harmful to animals living in the wild, causing them pain and in some instances death. Some examples are listed below. 

Using eDNA sampling, scientists found a strong positive correlation between the density of DNA belonging to ciliate protozoan, a type of parasite, and the mortality of barramundi, a type of fish.46 eDNA sampling, using the feces of wild rats, was found to be quicker, easier and far more sensitive than traditional methods, finding more varieties of parasites.47 eDNA sampling identified geographical areas where there were higher levels of Blastocystis, a parasite known to infect both humans and nonhuman animals.48 Helping to understand levels of parasitic disease in different geographical areas could be very helpful when trying to understand which location to address first. Lastly, four species of Cryptosporidium, a parasite known to infect both cows and human beings, were detected simultaneously by sampling cow feces.49

Species Detection

One of the most distinct advantages of eDNA sampling is that it is exceptionally good at detecting the presence of species. Some interesting examples are given below. 

When compared to camera trapping, eDNA sampling was able to detect far more small species including mouse-eared bats, deer mice, voles, and brown rats that would have normally eluded camera traps.50 eDNA sampling detected 44% more shark species than normal traditional underwater censuses, even some previously unobserved shark species in the study area.51 To detect particular animals, a team collected water samples from ponds and confirmed the presence of several land animals including the common raccoon, Norway rat, and the house mouse. The team described traditional surveying methods as laborious compared to eDNA sampling.52

How eDNA could help animals living in the wild

Currently eDNA sampling is primarily being used for conservationist purposes, and it would be encouraging to see eDNA sampling used to help individual animals instead. The following are some speculative examples of how eDNA sampling could be used to help animals living in the wild. 

Detecting disease in the future

While eDNA sampling is being used to detect disease, it is mostly being used in situations where animals are kept captive or to maintain a population of animals in the wild that are at risk of becoming extinct. However, eDNA can be used to help individual animals living in the wild. For example, animals living in urban areas are prone to diseases such as rabies. Rabies in bats has previously been detected using eDNA sampling in the form of feces53 and this could be an effective way to screen for the early detection of rabies in urban areas. It would be helpful to use eDNA sampling to test for these diseases in populations of animals living in the wild.

eDNA sampling could also be used to detect levels of parasitic DNA. Using existing research, there is the potential to understand in which geographical areas parasites are more likely to attack animals in the wild and these areas could be screened using eDNA sampling. Animals can suffer greatly due to parasitic diseases and early detection would make it more likely to avoid higher levels of suffering and mortality due to them.

Using eDNA to source DNA for analyzing biomarkers 

Biological age biomarkers act as indications as to whether an animal has experienced significant negative or positive life events during their lifespan.54 

At this time there is not one single biomarker that can be applied to all animals. However, it is likely that biological age biomarkers will be better understood in the future, and human beings will be able to identify life events that are particularly stressful for individual animals. Using biological age markers, once particular life events are found to cause serious psychological or physiological trauma to animals, it will be easier to understand effective ways to help them. Examples of negative life events could include fighting with another animal or suffering from starvation for a number of days.

It has been suggested that the length of telomeres (the ends of chromosomes) shortens as the biological age of vertebrates increases and the shortening process is sped up by negative life experiences.55 Analyzing the length of telomeres could be a potential way to understand the ways and reasons that vertebrates living in the wild suffer.

DNA samples taken for the purposes of analyzing telomere length are almost always taken using invasive techniques in the form of blood and saliva samples.56 While blood sampling will probably always be invasive, there are noninvasive methods to collect samples of saliva. One study highlighted the ease of eDNA sampling by collecting gnawed wood which contained the saliva of aye-ayes.57 Therefore eDNA sampling may be able to provide a noninvasive alternative to the collecting of samples to analyze telomere length and consequently to better understand the lives and suffering of animals living in the wild. 

Once there is sufficient evidence that a particular life event or environment could be stressful for an animal, then it may be possible to help her. Examples of ways may include moving naturally solitary fishes to environments with smaller populations of fishes thereby taking them out of a stressful environment,58 or rescuing mice from environments that have ultrasound frequencies which could cause them to exhibit depression-like symptoms.59

Detecting species

Species detection is a tool that is not currently being utilized in an obvious way that would help individual animals. For example, species detection is being used to understand whether a species is abundant or not in a particular area, but not to discover whether an animal is present in what could be an unsafe environment and in need of help. 

Being able to detect a particular species could be used in certain situations where animals could be in unhealthy or dangerous environments. If it is known where animals are prone to being trapped in urban areas, or environments that are toxic to animals, it would be helpful to use eDNA sampling as a tool to understand whether an individual is likely in danger. It can only be speculated at present how species detection would be used to help individual animals living in the wild. However, it offers a number of opportunities to be explored.

Conclusion

This article has focused on eDNA sampling and the ways in which it could potentially help to reduce the suffering of animals living in the wild. 

Right now, there are billions of animals suffering in the wild for a number of reasons, and there are many ways in which eDNA sampling can help these animals. These include using eDNA sampling instead of invasive trapping and handling methods in research whenever possible. In many instances eDNA sampling has been shown to be as accurate, more cost-effective and to require lower sampling effort than some invasive methods. eDNA sampling can also be applied in a number of environments which would prove to be trickier for invasive techniques. 

eDNA sampling can detect diseases in animals earlier than obvious symptoms may appear. Promoting the use of eDNA sampling for the early detection of diseases could bring attention to the plight of animals in the wild due to disease and lead to the development of vaccines and other treatments.

The situation of animals living in the wild is yet to be properly understood and more research is needed.  Currently eDNA sampling is more commonly used for conservationist purposes, for example, to help detect a species. eDNA sampling deserves more attention for its potential to help reduce the suffering of animals living in the wild. It can be used more extensively in ways that we already know are useful, such as providing data that allow us to know when to address diseases and stressful environments. There will be other applications that have not yet been explored or even conceived of yet. The technique is progressive and there is hope that it could revolutionize the ability to help animals in the near future. 

Notes

  1. Taberlet, P.; Waits, L. P. & Luikart, G. (1999) “Noninvasive genetic sampling: Look before you leap”, Trends in Ecology & Evolution, 14(8), pp. 323-327.
  2. Gagneux, P.; Boesch, C. & Woodruff, D. S. (1997) “Microsatellite scoring errors associated with noninvasive genotyping based on nuclear DNA amplified from shed hair”, Molecular Ecology, 6, pp. 861-868.
  3. Peters, C.; Nelson, H.; Rusk, B. & Muir, A. (2019) A novel method to optimise the utility of underused moulted plumulaceous feather samples for genetic analysis in bird conservation”, Conservation Genetics Resources [accessed 17 June 2020].
  4. Bradley, B. J.; Doran-Sheehy, D. M. & Vigilant, L. (2007) “Potential for female kin associations in wild western gorillas despite female dispersal”, Proceedings of The Royal Society B, 274, pp. 2179-2185.
  5. Kawai, K.; Shimizu, M.; Hughes, R. N. & Takenaka, O. (2004) “A non-invasive technique for obtaining DNA from marine intertidal snails”, Journal of the Marine Biological Association of the United Kingdom, 84, pp. 773-774.
  6. Lanyon, C. V.; Rushton, S. P.; O’Donnell, A. G.; Goodfellow, M.; Ward, A. C.; Petrie, M.; Jensen, S. P.; Gosling, L. M. & Penn, D. J. (2007) “Murine scent mark microbial communities are genetically determined”, FEMS Microbiology Ecology, 59, pp. 576-583.
  7. Franklin, T. W.; McKelvey, K. S.; Golding, J. D.; Mason, D. H.; Dysthe, J. C.; Pilgrim, K. L.; Squires, J. R.; Aubry, K. B.; Long, R. A.; Greaves, S. E.; Raley, C. M.; Jackson, S.; MacKay, P.; Lisbon, J.; Sauder, J. D.; Pruss, M. T.; Heffington, D. & Schwartz, M. K. (2019) “Using environmental DNA methods to improve winter surveys for rare carnivores: DNA from snow and improved noninvasive techniques”, Biological Conservation, 229, pp. 50-58.
  8. Lacoursiere-Roussel, A.; Côte, G.; Leclerc, V. & Bernatchez, L. (2016) “Quantifying relative fish abundance with eDNA: A promising tool for fisheries management”, Journal of Applied Ecology, 53, pp. 1148-1157.
  9. Deagle, B. E.; Chiaradia, A.; McInnes, J. & Jarman, S. N. (2010) “Pyrosequencing faecal DNA to determine diet of little penguins: Is what goes in what comes out?”, Conservation Genetics, 11, pp. 2039-2048.
  10. Ng, T. F. F. Kondov, N. O.; Deng, X.; Van Eenennaam, A.; Neibergs, H. L. & Delwart, E. (2015) “A metagenomics and case-control study to identify viruses associated with bovine respiratory disease”, Journal of Virology, 89 (10), pp. 5340-5349.
  11. MacArthur, R. A.; Geist, V. & Johnston, R. H. (1986) “Cardiac responses of bighorn sheep to trapping and radio instrumentation”, Canadian Journal of Zoology, 64, pp. 1197-1120.
  12. Bosson, C.; Islam, Z. & Boonstra, R. (2012) “The impact of live trapping and trap model on the stress profiles of North American red squirrels”, Journal of Zoology, 288(3), pp. 159-169.
  13. Lynn, S.E. & Porter, A. J. (2008) “Trapping initiates stress response in breeding and non-breeding house sparrows Passer domesticus: Implications for using unmonitored traps in field studies”, Journal of Avian Biology, 39, pp. 87-94.
  14. Fletcher, Q. E. & Boonstra, R. (2006) “Impact of live trapping on the stress response of the meadow vole (Microtus pennsylvanicus)”, Journal of Zoology, 270, pp. 473-478.
  15. Balcombe, J. P.; Barnard, N. D. & Sandusky, C. (2004) “Laboratory routines cause animal stress”, Journal of the American Association for Laboratory Animal Science, 43(6), pp. 42-51.
  16. Çelik, M. Y.; Duman, M. B.; Sarıipek, M.; Uzun Gören, G.; Kaya Öztürk, D. & Sedat Karayücel, S. (2018) “Growth and mortality rates of Cornu aspersum: Organic snail culture system, Black Sea region”, Journal of Agricultural Sciences, 25, pp. 189-196.
  17. Shonfield, J.; Do, R.; Brooks, R. J. & McAdam, A. G. (2013) “Reducing accidental shrew mortality associated with small-mammal livetrapping: An inter- and intrastudy analysis”, Journal of Mammalogy, 94 (4), pp. 745-753.
  18. Takacs, V.; Zduniak, P.; Panek, M. & Tryjanowski, P. (2009) “Does handling reduce the winter body mass of the European hare?”, Central European Journal of Biology, 4(3), pp. 427-433.
  19. Deguchi, T.; Suryan, R. M. & Ozaki, K. (2014) “Muscle damage and behavioral consequences from prolonged handling of albatross chicks for transmitter attachment”, The Journal of Wildlife Management, 78(7), pp. 1302-1309.
  20. Ponjoan, A.; Bota, G.; Garcia De La Morena, E. L.; Morales, M. B.; Wolff, A.; Marco, I. & Mañosa, S. (2008) “Adverse effects of capture and handling little bustard”, Journal of Wildlife Management, 72 (1), pp. 315-319.
  21. Field, K. A.; Paquet, P. C.; Artelle, K.; Proulx, G.; Brook, R. K. & Darimont, C. T. (2019) “Publication reform to safeguard wildlife from researcher harm”, PLoS Biol, 17(4) [accessed 17 June 2020]
  22. Piggott, M. P. (2004) “Effect of sample age and season of collection on the reliability of microsatellite genotyping of faecal DNA”, Wildlife Research, 31, pp. 485-493.
  23. Dell’Anno, A. & Corinaldesi, C. (2004) “Degradation and turnover of extracellular DNA in marine sediments: Ecological and methodological considerations”, Applied and Environmental Microbiology, 70(7), pp. 4384-4386.
  24. Dejean, T.; Valentini, A.; Duparc, A.; Pellier-Cuit, S.; Pompanon,F.; Taberlet, P. & Miaud, C. (2011) “Persistence of environmental DNA in freshwater ecosystems”, PLoS ONE, 6(8), [accessed 17 June 2020]
  25. Dell’Anno, A. & Corinaldesi, C. (2004) “Degradation and turnover of extracellular DNA in marine sediments: Ecological and methodological considerations”, op. cit.
  26. Leempoel, K.; Hebert, T. & Hadly, E. A. (2019) “A comparison of eDNA to camera trapping for assessment of terrestrial mammal diversity”, Proceedings of the Royal Society B, 287, [accessed 17 June 2020].
  27. Goldberg, C. S.; Turner, C. R.; Deiner, K.; Klymus K. E.; Thomsen, P. F.; Murphy, M. A.; Spear, S. F.; McKee, A.; Oyler-McCance, S. J.; Cornman, R. S.; Laramie, M. B.; Mahon, A. R.; Lance, R. F.; Pilliod, D. S.; Strickler, K. M.; Waits, L. P.; Fremier, A. K.; Takahara, T.; Herder, J. E. & Taberlet, P. (2016) “Critical considerations for the application of environmental DNA methods to detect aquatic species”, Methods in Ecology and Evolution, 7, pp. 1299-1307.
  28. Miaud, C.; Arnal, V.; Poulain, M.; Valentini, A. & Dejean, T. (2019) “eDNA increases the detectability of Ranavirus infection in an alpine amphibian population”, op. cit.
  29. Ibid.
  30. Shaw, J. L. A.; Clarke, L. J.; Wedderburn, S. D.; Barnes, T. C.; Weyrich, L. S. & Cooper, A. (2016) “Comparison of environmental DNA metabarcoding and conventional fish survey methods in a river system”, Biological Conservation, 197, pp. 131-138.
  31. Valentin, R. E.; Fonseca, D. M.; Nielsen, A. L.; Leskey, T. C. & Lockwood, J. L. (2018) “Early detection of invasive exotic insect infestations using eDNA from crop surfaces”, Frontiers in Ecology and the Environment, 16(5), pp. 265-270.
  32. Evans, N. T.; Shirey, P. D.; Wieringa, J. G.; Mahon, A. R. & Lamberti, G. A. (2017) “Comparative cost and effort of fish distribution detection via environmental DNA analysis and electrofishing”, Fisheries, 42(2), pp. 90-99.
  33. 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), pp. 4023-4032.
  34. Ibid.
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