Download ethical guidelines for field research on vertebrates animals

CHARLES DARWIN UNIVERSITY
GUIDELINES FOR FIELD RESEARCH ON VERTEBRATES
About this document
These guidelines have been prepared by the Charles Darwin University Animal Ethics Committee (AEC) with
the objective of providing guidance to field researchers handling native vertebrate animals in the field in the
Northern Territory. The guidelines do not cover laboratory experiments using native wildlife. The AEC can be
consulted for further advice on issues considered below, and would be pleased to receive suggestions for
further development or refinement of the guidelines.
1. Introduction
The conservation of wildlife depends on an adequate knowledge of their status, distribution and ecology. Field
research contributes to such knowledge, and is a vital element in the management of wildlife, particularly of
threatened, exploited and pest species. However, field research is potentially intrusive, disturbing and/or
destructive to individual animals and possibly to whole populations of animals. These guidelines aim to ensure
that researchers are aware of the potentially detrimental effects of their actions and of the means of minimising
negative impacts.
These guidelines complement the recent (2004) 7th edition of the Australian code of practice for the care and
use of animals for scientific purposes prepared by a joint working party of NHMRC, CSIRO, ARMCANZ and the
state and territory governments. The Code of Practice contains an introduction to the ethical use of animals in
field research and should be referred to for broader issues. These guidelines offer detail specific to research
on wildlife in the Northern Territory. Other sources for more detail include Tribe and Spielman (1996) for the
restraint and handling of captive wildlife, Oring et al. (1988) for a detailed review of the use of wild birds in
research, Cuthill (1991) on the ethics of animal behaviour studies, ASIH et al. (1987a, b) for detailed guidelines
on research on fish, amphibians and reptiles, and NHMRC (1995) on the care of individual Australian native
mammals.
The trapping of animals for food or other products has been a universal feature of human ecology for
thousands of years. Many of the techniques now used by wildlife biologists are based on methods developed
by hunters, and require considerable modification to ensure that the wellbeing rather than the destruction of the
animal is given primary consideration. Many other field techniques (such as radio-tracking) have developed
rapidly over the last few decades, and animal welfare issues have sometimes lagged behind advances in
technological sophistication. Sometimes, adverse impacts are difficult to predict or detect. In many cases,
although separate impacts may be small, a series of effects on the same subject may produce more
substantial cumulative impact.
Carefully selected experimental design will often reduce the impacts of wildlife research. In many cases, the
amount of data required to answer the experimental questions can be determined a priori by statistical power
analysis. In some cases, judicious use of modelling may signal the most efficient modes of data collection.
This document considers the more common elements of field research dealing with native animals, and
outlines the risks to the subjects and preferred techniques for minimising such risks. A more comprehensive
treatment is beyond its scope, as the research aims of field biologists and the techniques now used, are so
extraordinarily varied.
2. Guiding principles
There are a few universally accepted guiding principles for field research against which research objectives
and methods should be measured. These are:
 to minimise the negative impacts of field research on wildlife individuals and populations;
 to treat animals used in field research with as much care as is expected for the use of animals in
laboratory-based research. Specifically, to use humane methods in all aspects of capturing, handling,
holding and releasing animals;
 the techniques of field research should alter the habitat as little as possible;
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Guidelines for field research on vertebrates
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proposals for field research should demonstrate that researchers have adequately considered animal
ethics issues, and made every attempt to minimise their negative effects;
researchers should aim to minimise the disruption to normal activities of individual animals;
the negative impacts of research on some individual animals should be weighed against the likelihood of
beneficial outcomes of the research for the species (or environment) as a whole;
researchers should be suitably experienced and competent in the procedures being implemented;
projects should have feedback mechanisms built in to monitor the impacts on the animals affected, with
action taken where necessary, to alter procedures to reduce such impacts;
researchers should consider possible wider impacts of their research, notably on non-target animals;
ethical guidelines should apply to feral and other “unwanted” animals: e.g., individual feral animals should
not be exposed to any more suffering than native animals;
impacts should be documented honestly, and this information should be reported in order to enable
refinement or a more informed assessment of techniques used or proposed in subsequent studies.
3. Observation and passive recording
Much field research can be undertaken without the need to trap, handle or mark individual animals.
Axiomatically the detrimental impacts of such research are likely to be less than research using more intrusive
techniques. However, the mere presence of researchers may lead to a failure to nest, increased risks of
predation, increased stress and disturbance to normal behaviours. This applies especially to large nesting
colonies, to low-level aerial surveys, to flighty species, to the use of attractants to promote detectability and/or
observability (e.g. decoys, play-backs of territorial calls) and to frequent site visits. Individuals of rarely-seen
bird species are known to have suffered harassment by, and ultimately death through, the enthusiasm of birdwatchers.
Recent technological advances have allowed the development of recording devices for detecting the speciesspecific calls of many animals (notably frogs and bats), and hence the assessment of species composition and
abundance without handling animals or any other interference. The use of such devices is obviously less
intrusive than traditional methods and is to be encouraged wherever possible, but the novelty of these
techniques means that many recorded calls still need verification by taking voucher specimens, and many
research questions cannot be addressed so simply.
A fundamental guideline to research involving observation is the minimisation of disturbance and interruption to
the “normal” behaviour of wild animals. This can be achieved by the observer maintaining a sufficient distance
from the subject animals so as to be non-threatening; by the use of “hides”; by minimising the use of
attractants or other artificial stimulants; by minimising the time spent in close contact; and by restricting
observation to times at which disturbance is least likely to have impact (e.g. avoiding visiting seabird colonies in
the middle of the day). Further information on measures to reduce impacts of visits to seabird colonies are
given in WBM Oceanics Australia and Gordon Claridge (1997).
4. Capture
The capture of wild animals is often necessary for ecological investigations. There are a number of preferred
techniques which minimise the risks associated with capture. The Code of Practice notes that the over-riding
principle on trapping protocol is to minimise the impact on both target and non-target species, and that
researchers should consider the time the animals will spend in the traps; protection of trapped animals from
predators, parasites and/or disease (e.g. through ensuring that traps are clean); protection of trapped animals
from environmental effects such as dehydration, hypothermia and drowning; deprivation of food and water;
potential for impact via disruption of social structure; potential for impact on dependent young; and trap design
and use (including deactivation after the research period, appropriate size and construction).
4.1. Capture by hand
The most common technique for capturing amphibian, reptilian and some mammalian species is by active
search and hand-capture. This involves a search for the sites that animals roost or shelter in and their removal
from under logs or rocks, behind loose bark, in old bird nests, or in holes and burrows. In some cases, this
process may disturb or destroy the shelter site, and hence decrease the survival chances of the animal.
Researchers should aim to minimise damage to important shelter sites, and wherever possible repair such
damage.
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4.2. Terrestrial mammal traps
A wide range of traps are commercially available for capturing mammals. Until recently, break-back traps
(such as typical household mouse-traps) were widely used in wildlife surveys. There is no longer any
justification for the use of destructive traps in wildlife research. Most commonly-used live mammal traps are
either variants on wire-mesh cages or enclosed aluminium boxes, with trip/trapdoor mechanisms. Both may
have substantial impacts on the animals they catch. Aluminium traps (such as Elliot traps) may become very
cold or very hot. Cage traps may better reflect ambient temperatures, but many animals may injure their heads
by poking their noses through the mesh. Both sorts of traps may occasionally injure tails when trapdoors shut.
Ants may be attracted to trap bait, and cause discomfit or even death to trapped animals. Some types of
mammals (e.g. quolls and bandicoots) may be especially stressed by trapping.
Trap impacts should be reduced by:
 ensuring that all traps are clearly laid out and marked, so that none are missed and left behind;
 ensuring that all traps are out of direct sunlight, and that they are checked within one hour of dawn - this is
achieved by balancing the number of people, number of traps laid out, the time researchers get up and the
distance between traps;
 ensuring that the amount of bait in traps is sufficient to provide an adequate food resource for trapped
animals;
 placing insulation material within (e.g. grass, old cloth) or around (plastic bags) traps when overnight
temperatures may become very low, and/or when rain is possible;
 ensuring the trap will not be submerged in the event of rain;
 closing traps on very cold nights and during hot days; and
 monitoring ant activity around traps, and spraying insecticides if ant activity is high.
4.3. Pitfall traps
Pitfall traps are plastic or metal tubes or buckets dug into the ground. Depending upon their size, a variety of
animals fall into them, and some of these cannot then get out. The probability of catching animals is often
increased by the use of driftline fencing to direct animals into the pit. Pitfall traps may have a number of
adverse impacts on the animals they catch:
 trapped animals may be damaged or consumed by other trapped animals (particularly ants, but also
centipedes, beetles, spiders) or by animals which can access the traps without themselves being caught
(e.g. snakes, goannas and dasyurids);
 pitfall traps may be very exposed to weather, becoming very hot on warm days, and potentially flooded with
rain;
 animals caught in pitfalls may be deprived of their food requirements;
 lactating females with dependent young may be caught in the traps.
As with terrestrial mammal traps, these effects can be minimised by:
 frequent checking (2-4 times a day, depending upon weather);
 ensuring that all traps are checked within one hour of dawn;
 placing adequate shelter and insulation in the bottom of pit-traps (a wet Chux-cloth or equivalent may be
the best material, as it prevents dehydration in frogs - but soil and leaf litter is generally OK);
 placing a small hole in the base of the bucket to allow rainfall to drain (though this will not work in heavy
rainfall, and can let groundwater in);
 monitoring ant activity around traps, and spraying insecticides if ant activity is high;
 covering with a raised lid that still allows animals to fall in;
 examining and determining the lactation status of the captured animal and releasing immediately any
lactating females.
Some museum collectors and researchers place a layer of killing/preservative fluid in the bottom of pits (”wet
pitfalls”). This technique obviously has a very high casualty rate, including many non-target animals, and
should not be used.
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4.4. Traps for aquatic vertebrates
Baited drum traps can be used to catch freshwater turtles. These typically are cylindrical in shape and
composed of netting on a metal frame with a funnel entrance on each end and bait suspended about mid
chamber. Like many fish traps the trap retains animals that are unable to locate the inverted funnel exit.
Collapsible drum traps (Legler 1960) and variations on the drum trap that reduce the chance of escape
(Kennett 1992) have been used successfully. Often these traps are set below the surface and traps must
be checked regularly (say every 1-1.5 hours depending on conditions and species to prevent drowning.
Alternatively traps can be set at the surface by fixing them to riverside vegetation or enclosing floats. In
some areas this doe not reduce capture rates. All traps must be retrieved or they will continue to trap
turtles after the researcher has left a site. Other factors to consider are the rare capture of crocodiles and
other fauna such as goannas. More regular checking may be required in situations where non-turtle
aquatic reptiles are likely to be caught.
4.5. Nets for catching birds and bats
Mist nets are routinely used for the capture of birds and bats, and work by entangling flying animals in a very
fine mesh. Damage to captured animals may arise through injuries sustained when hitting the coarser shelf
strings, predation (typically by ants, other birds or snakes), exposure (to excessive heat or cold) and/or from
poor handling during untangling. Such risks have been well documented and mechanisms for minimising them
well considered (e.g. Wilson et al. 1965; Lowe 1989), not least through a rigorous licensing system dependent
upon substantial training and supervision by experts. As general principles, nets should not be operated at hot,
cold or rainy times; the number of qualified researchers should be appropriate for the maximum number of
animals which may be expected to be caught; nets should be checked at intervals of no more than 10 minutes,
and birds should be removed from nets within 10 minutes of capture.
Cannon nets have been used more recently to trap flocks of birds (typically waders). These operate by firing a
net strung between projectiles. As with mist nets, there is a rigorous training and licensing process required
before researchers are permitted to use these devices. The use of cannon nets should be guided by similar
principles to that described above for mist nets, with extra concerns relating to care with the firing of heavy
projectiles around and above birds (a problem which becomes more serious if birds take off at the time of
firing), and the timing and positioning of firing in relation to tides (for shorebirds). A more rudimentary analogue
of cannon nets, “clap trap” (small nets attached to poles which snap together when released from guys), has
been used recently to capture finches, but has proven to have an unacceptably high rate of mortality.
4.6. Harp traps
Harp traps (Tidemann and Woodside 1978) are now routinely used for capturing bats. These consist of a bank
of vertical fishing-lines leading to a holding bag. Flying bats hit the lines and fall into the bag, from which they
are extracted. Harp traps should be checked through the night at 3-4 hour intervals. Mortality or injury directly
attributable to the trap is generally recognised to be very low, but cases have been reported of cannibalism of
bats within the holding bag, or small predators (e.g. snakes, dasyurids, rodents) entering the bag and
consuming the captured bats. The entry of non-bats can be minimised by careful trap placement (e.g.
ensuring no vegetation leads from the ground to traps). Traps can also be checked during the night to reduce
over-crowding and other interactions between captured bats. Traps should not be set on very cold, very hot or
very rainy nights.
4.7. Capture of larger terrestrial mammals
A variety of methods are used to catch macropods (for a summary see Coulson 1996) and the suitability of
a particular technique depends on the species and situation.
Smaller macropods can be caught using baited traps (Pollock and Montague 1991) or drive fences
(Vernes 1993). They can also be caught in trap yards and either herded into nets (Lentle et al. 1997) or in
some cases caught with hand nets. Early attempts to catch larger macropods involved trap yards and
physical handling of the animals. This resulted in considerable mortality in some studies (e.g. Keep and
Fox 1971). Other methods for catching medium and large macropods include “stunning” (Robertson and
Gepp 1982), cannon netting (Clancy and Croft 1992), bait drugs such as alpha-chloralose (Arnold et al.
1986), draw string traps that exploit movement through fences (Coulson 1997) and darting (Higginbottom
1989; Stirrat 1997).
Prevention of physical injury during the capture process should be a priority and this usually depends on
the skill of the worker involved. However, emphasis in all of these techniques should also be on prevention
of capture myopathy, a potentially serious side effect of the capture process that can cause high mortality.
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Capture myopathy, or capture stress, is a disease associated with the capture and handling process, the
main feature being degeneration of skeletal and/or cardiac muscle which causes physiological problems
(Shepherd et al. 1988). In acute cases the animal may collapse and die immediately, but death may occur
weeks after capture (Shepherd 1984). In non lethal situations capture myopathy may result in debility or
impairment of normal function, predisposing the animal to predation or other environmental stresses.
Capture myopathy can occur in macropods of all sizes (Shepherd 1983).
Onset of capture myopathy requires only psychological stress in addition to some exercise, so it may
develop soon after a capture process has started and long before an animal is actually handled. Onset
can be reduced by avoiding prolonged exercise during capture (e.g. chasing), by blindfolding the animal
and placing it in a dark area, and by restricting free movement to avoid rupture of necrotic muscle
(Shepherd 1984). To reduce trauma a tranquilizer or sedative can be administered at the time of capture.
Advice should be sought from a wildlife veterinarian in choosing a suitable drug which will achieve the
desired results. Capture myopathy can proceed while a drug takes effect, and some drugs may immobilise
an animal but have no tranquilizing properties. Darting, or remote injection of a tranquilizer, is not suitable
in all situations but has the potential to greatly reduce the likelihood of capture myopathy.
4.8. Capture on and around nests and/or roosts
Many animals are easiest to catch at sites which they must visit, such as nests or communal roosts. Such sites
present researchers with relatively easy opportunities to catch particular individuals or a number of specimens.
However, interference, such as by trapping, at these sites may pose a substantial risk of adverse impacts. The
possibility of disruption to an animal’s essential activities should be carefully considered in such studies, and
alternatives to these home invasions should be explored first. For example, Helman and Churchill (1986)
recommend that cave-roosting bats should be investigated primarily by counts at exits rather than by the
researchers entering the caves.
4.9. Other trapping methods
There are very many other approaches for trapping animals, of very variable ethical acceptability. Helman and
Churchill (1986) list many for bats, and McClure (1966) lists many for birds. Researchers should be able to
justify using non-standard trapping techniques, and be prepared to adapt such techniques rapidly in light of
their experiences with their initial use.
Nesting sea turtles can be readily examined as they haul up the beach to lay eggs but this provides no
information on juvenile, non-nesting or male turtles. These must be caught in the water. In some locations
turtles can be guided into an enclosure by fixed nets or may become trapped in rock pools on tidal rock
shelves. In other areas the most common method is the turtle rodeo where the turtles are searched for
then followed by motor boat until the diver can enter the water and grab the shell of the turtle and guide it
to the surface. Where visibility is good and water is relatively shallow, capture success is improved by
following the turtle to it begins to tire then securing it.
5. Handling and holding animals
The Code of Practice offers guidelines for the handling of wild animals. The over-arching principle is that
captured animals should be handled in a way that minimises the risk of injury or stress-induced disease. The
Code of Practice suggests that this can be best achieved through firm and quiet handling; keeping handling
and restraint time to the minimum needed to achieve the scientific or educational objectives; using sufficient
competent persons to restrain animals and prevent injury; using techniques and timing appropriate to the
species; and using, where appropriate, chemical restraint if animals are to be held for more than a short time.
Handling techniques should also consider the desirability of minimising damage to the handler – bitten or
scratched researchers may be more likely to stress or injure animals than those in better control of the
situation!
Captured animals may be retained for identification, marking and measurement before release. In general, the
shorter this time the better, although in the case of nocturnal species, it generally better to release individuals
at nightfall. Careful consideration should be given to whether it is necessary to retain animals for any length of
time, as some animals (e.g., some species of bats) are easily stressed and may die in captivity.
During the holding time animals should be kept in containers which provide them with a comfortable
temperature, cannot lead to injury, and provide adequate ventilation. It may be appropriate to provide food and
water. As a rule of thumb, mammals and reptiles are best held in cloth bags, frogs in plastic bags with some
water, and birds in either cloth bags or holding cages. Held animals should be monitored frequently for signs
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of distress, although this needs to be balanced against the desirability of limiting disturbance. Containers
should house only single animals at any time. Containers should be cleaned frequently to minimise chances of
spread of parasites and diseases.
Animals should be released near the point of capture, at a time consistent with the species’ normal activity
rhythm. Researchers should consider the possibility that, during holding, animals may be exposed to disease
or parasites, hence there may be some risk of the researcher inadvertently threatening that population. The
hygiene practised by the researcher, whether the retained animal is transported and the length of time the
animal is retained, are all likely to affect the risk of exposure to novel pathogens.
6. Tagging, marking and banding
The ability to identify individual animals is a key requirement for many wildlife studies. In some instances this
can be done non-invasively through documentation of idiosyncratic markings, however, in the majority of
cases, researchers needing to recognise individuals must apply some identifying mark. A wide variety of
recognition aids have been used, including painting, tags, leg-bands, neck-bands, streamers, ear clipping, toe
clipping, shell clipping, transponders, and branding. The Code of Practice offers the general rider that the
technique used should be that which causes the least distress within the context of the research proposal and
the least interference with the normal functioning of the animal.
Lowe (1989) and subsequent information from the Australian Bird & Bat Banding Scheme (GPO Box 8,
Canberra, ACT, 2601) provide detailed recommendations on marking schemes appropriate for Australian birds
and bats. They note that there may be substantial variation between species in their response to tagging (e.g.
many parrots chew leg bands and can then damage their legs unless the band is made of particularly strong
metal). Some tagging schemes have been shown to increase predation (e.g. patagial tags on cockatoos) or to
produce a high risk of subsequent injury (e.g. wing bands on many bats), and their use is now prohibited or
discouraged. In some bird species, colour bands have been demonstrated to alter breeding success and other
social parameters.
7. Radio-telemetry
Researchers should refer to Kenward (1987) for a wide-ranging discussion of techniques of radiotelemetry. Of
particular interest is Chapter 5, "Tag Attachment" which discusses topics such as avoiding adverse effects,
attachment techniques and detachment. Investigators new to the techniques of transmitter attachment should
also refer to literature related to the group of animals they are working with, and the internet for the latest
developments in telemetry techniques.
In the past, emphasis has been placed on the percentage weight of the transmitter package as compared to
the weight of the animal. For example a rule of thumb for birds is 5% while for some terrestrial animals may
comfortably carry 15%. However, investigators should not think that this is the only important factor.
Transmitter profile may be more important than weight for some animals. Investigators need to consider the
life style and habits of the animals under study. For example, animals inhabiting small crevices may become
wedged if the transmitter protrudes from the line of the body regardless of transmitter mass. If possible, it may
be advantageous to attach (or implant) the transmitter on an individual in captivity and observe the animal for
signs of impaired movement, irritation or rubbing caused by the transmitter.
There are a number of modes of attachment including harnesses, glue, collars and implantation. There is no
one best method: it depends on the animal and its habits. The guiding principle is to minimise the impact on
the movement of the animal and to avoid short-term and long-term injury resulting from the transmitters. In
some cases it is possible to design harnesses or other attachment materials so that they have a weak link,
ensuring that the transmitter is detached if it becomes entangled. In these cases, and with the use of glue, it is
likely that the transmitter will eventually become detached in the field. Otherwise, it is desirable to recapture
the animals (before the transmitter fails) and remove the transmitters. However, it is always possible that the
transmitter will fail or the animal may move a long distance, making recapture unlikely. For this reason, it is
desirable to design transmitters and their attachment with this possibility in mind so that long-term attachment
will not result in injury.
8. Collecting specimens and body part samples
The deliberate killing of wild animals for scientific collection remains a contentious issue. In places such as the
Northern Territory, where many groups have been poorly inventoried or remain taxonomically unresolved,
some collections may be required to confirm species’ identity or to provide the basis for the description of new
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taxa. The following guidelines for collection of specimens should be followed.
 Animals should be killed using appropriate euthanasia techniques. Reilly (1993) provides a detailed
description of recommended and acceptable procedures, a summary of which is included in the AEC
guidelines. Note that some of these techniques require considerable experience to be used appropriately.
 Maximum use should be made from collected specimens, to minimise the number of specimens required all specimens should be properly preserved and later deposited in museums. Wherever possible tissue
samples should be taken just after death and stored appropriately for possible subsequent genetic
analysis.
 Impacts upon the local population should be considered – the number of specimens taken should not have
a significant impact upon the viability of the remaining population.
 Collection of specimens should be undertaken only when non-destructive techniques (e.g. blood
sampling, hair analysis for specific identity) are inapplicable or impractical.
9. Dietary analysis
Diet is an important component of ecology, and hence its consideration is often a vital aspect of wildlife
research. Some methods of investigating diet are non-invasive (e.g. observations of foraging, faecal analysis):
other techniques are variably discomforting or destructive. There is now no justification for killing animals
solely in order to obtain stomach samples.
Stomach flushing (typically using saline solution) is now used routinely for many vertebrates and is generally
regarded as fairly benign, provided that the operator is competent with the procedure.
Probably more intrusive and risky is the use of chemical emetics. Many have been used for dietary studies on
birds, with notably patchy success and impacts.
Physical manipulation of food items in the crop (a transparent food storage sac just below the skin of the neck)
has been used for many granivorous birds, and low rates of impact are generally reported (e.g. Zann and
Straw 1984).
10. Measuring and sexing
A wide range of measurements are routinely taken on captured wild animals in order to assess condition, sex
and/or age, or for specific identification. Most are routine and non-invasive but special care should be taken
when measuring more fragile or sensitive animals or parts, such as the wings and ears of bats. Reardon and
Flavel (1987) provide a good guide for holding and measuring bats, as does Lowe (1989) for birds.
Snakes and lizards can be sexed by determining the presence or absence of hemipenes, which are paired
structures at the base of the tail of males. In some cases the hemipenes can be everted by gently rolling the
thumb from side to side just posterior to the cloacal opening. If there is eversion, care should be taken that the
structure returns to its internal position before the animal is released.
A second method often used to detect hemipenes is to probe for them using a blunt probe, or in the case of
small animals, a blunt piece of fishing line. After the probe is inserted into the lips of the cloaca, it can be
gently pushed toward the tip of the tail on either side of the midline of the animal. If the individual is a female
the probe will meet resistance immediately, but the probe will extend into the hemipenes of males. Anyone
unfamiliar with these techniques should have them demonstrated by an experienced person.
Many species have bones in the hemipenes and sex can be determined from radiographs.
In many groups of small bats, penile morphology is the most reliable taxonomic feature. Reardon and Flavel
(1987) provide simple instructions for inspection in live animals.
11. Reproductive studies
Reproduction is a fundamental component of the biology of wild animals and hence of valid interest to
researchers. However, investigation of reproduction may be a field particularly prone to detrimental impacts
from the observer. The visits of researchers to nests, maternity roosts or other reproductive sites may lead to
increased predation, short-term or permanent abandonment by parents, damage to eggs or dependent young,
or premature departure of young. Impacts may be magnified where reproduction is concentrated in colonies.
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For reproductive studies of birds, Oring et al. (1988) recommend minimising visits and/or impacts through the
use of telescopes, hides, careful timing of visits, and gradual habituation. Helman and Churchill (1986)
suggested that maternity roosts of bats should not be disturbed.
The breeding biology of turtles and crocodiles has been the subject of much research in the Northern Territory.
Researcher impacts can be minimised through care taken to re-conceal nests after visits; to replace eggs in
their original position after inspection, and by minimising time spent at nests. Additionally, disruption to laying
female marine turtles should be kept to a minimum..
12. Habitat manipulation
Much ecological research in northern Australia considers the consequences of land management by simulating
different management regimes with experimental habitat manipulation. The landscape-scale fire experiments
carried out by CSIRO in Kakadu National Park are an example of this type of research. Some animals in the
research area are inevitably killed, or harmed by the reduced habitat quality following the experimental
treatment (e.g. Griffith’s and Christian 1996). The ethical costs of such research need to be balanced against
the likelihood of deriving results which can be translated to better management. Experiments involving
repeated destructive manipulation – beyond the bounds of that representative of real world management –
should be avoided.
References
American Society of Icthyologists and Herpetologists (ASIH), American Fisheries Society and American
Institute of Fisheries Research Biologists. (1987a). Guidelines for use of fishes in field research. Fisheries
(Bethesda) 13, 16-23.
American Society of Icthyologists and Herpetologists (ASIH), The Herpetologists’ League and The Society for
the Study of Amphibians and Reptiles. (1987b). Guidelines for use of live amphibians and reptiles in field
research. Gainsville, Florida.
Arnold, G.W., Steven, D., Weeldenburg, J. and Brown, O.E. (1986). The use of alpha-chloralose for the
repeated capture of western grey kangaroos, Macropus fuliginosus. Australian Wildlife Research, 13: 527533
Clancy, T.F. and Croft, D.B. (1992). Population dynamics of the common wallaroo (Macropus robustus
erubescens) in arid New South Wales. Wildlife Research, 19: 1-16.
Coulson, G. (1996). A safe and selective draw-string trap to capture kangaroos moving under fences.
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