Download guidelines for field research on vertebrates

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 (1997) 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 radiotracking) 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;
 proposals for field research should demonstrate that researchers have adequately considered animal
ethics issues, and made every attempt to minimise their negative effects;
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Guidelines for field research on vertebrates
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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 bird-watchers.
Recent technological advances have allowed the development of recording devices for detecting the
species-specific 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.
4.4. Traps for aquatic vertebrates
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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
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prevention of capture myopathy, a potentially serious side effect of the capture process that can cause
high mortality. 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
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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 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
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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 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
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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.
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.
Wildlife Research, 23: 621-627
Cuthill, I. (1991). Field experiments in animal behaviour: methods and ethics. Animal Behaviour 42,
1007-1014.
Griffiths, A.D. and Christian, K.A. (1996). The effects of fire on frillneck lizard, Chlamydosaurus kingii, in
northern Australia. Australian Journal of Ecology 21, 386-398.
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