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PLANNING A STUDY AND RECOMMENDED SAMPLING
A Brief Prepared by the Biological Survey of Canada
(Terrestrial Arthropods) 1994
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Knowledge of biodiversity is important for wise management and use of
the earths resources. Terrestrial arthropods (insects and their relatives) are by
far the most diverse groups of animals and important contributors to biodiversity.
However, a synopsis of techniques suitable for assessing diversity for terrestrial
arthropods is not readily available to many of those responsible for general assessments
of biodiversity. This brief therefore offers general guidelines for planning a study of
arthropod biodiversity, including attention to long-term planning, choice of taxonomic
groups, and the resources required for sampling, sorting and identification. The brief
recommends in some detail the specific sampling methods appropriate for this purpose. It
proposes a standard sampling protocol for the assessment of regional biodiversity,
suggesting that any such general inventory should include, at a minimum, Malaise,
flight-intercept and pan traps, as well as behavioural extractors such as Berlese funnels,
and it presents some estimates of the time required to process samples, for use in
planning a budget. The major current impediment to properly planned and executed studies
of arthropod diversity is the limited number of systematics experts available to identify
species. Resources for systematics support therefore should be included in project
PLANIFICATION DUNE ÉTUDE ET TECHNIQUES DÉCHANTILLONNAGE RECOMMANDÉES
La connaissance de la biodiversité est importante si
lon veut effectuer une gestion sage de lutilisation des ressources de la
terre. Les arthropodes terrestres (les insectes et leurs parents) constituent de loin les
groupes danimaux les plus diversifiés et contribuent grandement à la
biodiversité. Bon nombre des personnes responsables des évaluations générales de la
biodiversité ne disposent toutefois pas dun sommaire des techniques convenables
pour lévaluation de la diversité des arthropodes terrestres. Le présent précis
offre donc les grandes lignes pour la planification dune étude de la biodiversité
des arthropodes, dont une prise en considération de la planification à long terme, du
choix des groupes taxinomiques et des ressources requises pour léchantillonnage, le
tri et lidentification des arthropodes. Le précis recommande et décrit, en
détail, des méthodes déchantillonnage particulières convenant à cette fin. Il
propose un protocole déchantillonnage standard pour lévaluation de la
biodiversité régionale, mentionnant que tout inventaire général de ce genre devrait
comprendre, au moins, des pièges de Malaise, linterception darthropodes en
vol et des bacs jaunes, en plus dextracteurs éthologiques tels que
lextracteur de Berlese. De plus, il donne une certaine évaluation du temps requis
pour traiter les échantillonnages, ce qui peut faciliter la planification dun
budget. Le nombre limité dexperts systématiques pouvant identifier les espèces
constitue présentement le principal obstacle à la planification et lexécution
détudes sur la diversité des arthropodes. Il faudrait donc inclure dans les
budgets des projets des ressources pour lappui systématique.
Biodiversity has received recent national and international recognition.
The importance of biodiversity arises from the fact that the world
depends on self-sustaining biological systems that include many kinds of
organisms. Knowledge of biodiversity is required to understand the
natural world and the natural and artificial changes it may undergo; and
in turn, such knowledge permits the wise use and management of
ecosystems, both as elements of natural heritage and as reservoirs of
actual and potential resources.
biodiversity has been recognized as important in this general context,
not all of the methods for the actual study of diversity in particular
habitats are well known. In particular, standard methods are required to
assess the overwhelming numbers of insects, mites, spiders, and their
relatives, which form more than 75% of the world’s known species, and
present the greatest problems in arriving at estimates of regional
brief focusses on sampling methods appropriate to assess the taxonomic
diversity of terrestrial arthropods. It also emphasizes the importance
of proper long-term plans for any study of biodiversity. Carefully
developed plans are especially critical for arthropod inventories
because extensive and repeated sampling is required to capture the many
species with widely different habits, very large amounts of material are
generated, and it is difficult to identify many of the species.
Biological diversity, or biodiversity, has been used
to refer to almost any measure (taxonomic, numerical, genetic, etc.) of the variety of
organisms that live in a particular place. Although many different definitions of
biodiversity have been developed for particular uses, the focus here is on taxonomic
diversity or species richness the total number of kinds of organisms within a given
area, habitat, or community. Such assessments emphasize species, the functioning entities
in nature and the categories by which all biological information is organized and
retrieved. An emphasis on the methods required to obtain reliable measures of species
richness recognizes sampling and identification of species as the essential baseline for
understanding diversity: properly conducted inventories are the core of future endeavours.
Several types of analysis that use additional information, notably species abundance as
well as the number of species present, generate indexes of diversity of potential value
for understanding community structure (e.g. Pielou 1975; Kempton and Taylor 1976;
Southwood 1978; Magurran 1988). Such analyses are beyond the scope of this brief. However,
many of the techniques listed here are suitable for simultaneous assessments of taxonomic
and community diversity, especially by standardizing sampling effort.
Planning a study
It is important to emphasize at the outset that it is not now
possible, nor will it be possible in the foreseeable future, to inventory more than a
carefully chosen subset of the arthropod fauna of a region or habitat, because as many as
half of our insect species either are undescribed or cannot be identified at present.
Consequently, any study of arthropod biodiversity must be planned very carefully (Table
1). Rosenberg et al. (1979) provide a general discussion of Canadian arthropod surveys and
the elements of an ideal survey.
|Table 1: Components
of a properly planned biodiversity study
- Establish goals
- Select groups for study
- Decide how to deal with unnameable species
- Assemble financial resources for sampling and sorting
- Arrange for resources and systematics expertise for
- Define sampling methods
- Ensure follow-up (curation of voucher specimens, publication
of results, etc.)
The goals of a study dictate its plan. A general inventory requires substantially
different resources and sampling protocols than a more specific study. What are the best
means of trapping a diversity of predatory beetles? Will specimens be released following
their capture, requiring live-traps to be used? Will a systematist be willing to identify
moths taken from a fluid preservative? These and other questions must be anticipated and
dealt with early in the planning process. Sampling protocols might also be selected to
allow the results of a study to be compared in a standard way to previous studies or to
other ongoing studies.
It is not normally feasible to inventory all taxa of terrestrial arthropods. The choice of
taxa to be inventoried is governed mainly by the goals of the study, available resources
and systematics support. Several arthropod taxa can be identified as meeting criteria such
as habitat specificity, diversity, vagility, or other biological attributes relevant to
the goals of the study, but it will be necessary to choose practical candidates from that
list of potential taxa. Probably the best way to narrow the list is by assessing the
available systematics support (see below).
Because relatively few groups of terrestrial arthropods are well known, it will not be
possible to name many of the species collected, especially in diverse groups of small body
size. There are four ways to deal with these unnameable species:
- Taxa are sorted to morphospecies and assigned an identifying
number (e.g. Genus A sp. 1). This method allows species diversity to be assessed, but
requires a competent systematist to sort the taxa into morphospecies. It prevents
extraction of information on species biology from the literature, and prevents comparisons
with other studies unless the specimens from the other study have been similarly treated
by the same systematist.
- Taxa are treated at some higher taxonomic level (e.g. Genus A
spp.). This method is of limited value, because even though it requires accurate sorting,
preferably but not necessarily by a competent systematist, it masks significant
differences in natural history attributes of congeneric species, prevents detailed
analysis and prohibits detailed comparisons with other studies.
- Only taxa that can readily be identified are included. If
several taxa meet the criteria defined by the goals of the study, candidates can be
selected from that list of potential groups. As already noted, one of the best ways to
narrow the list of otherwise suitable potential candidate taxa is to look at the available
systematics support. What can be identified to the desired or required level? A few
groups, notably butterflies and large moths, dragonflies, and other groups covered by
comprehensive handbooks such as those published in the Agriculture Canada Insects
and Arachnids of Canada Handbook series, might be identifiable without direct
involvement by professional systematists. Such readily identifiable arthropod taxa remain
the exception and not the rule, and participation by professional systematists is nearly
always required. Although such a procedure makes a study with limited support feasible, it
does little to resolve the lack of knowledge about many insect groups.
- Resources are acquired to resolve systematics problems.
Biodiversity proposals that aim to solve rather than avoid the problems created by
arthropod biodiversity should include a budget item for support of professional or student
systematists to study the taxonomy of several of the key groups being sampled. This is the
optimum solution to the problem of unnameable species.
Resources for sampling and sorting
Because arthropod diversity is high and large numbers of specimens have to be processed,
any sampling programme involving arthropods is time consuming and expensive. Removal of
specimens from bulk samples, and the preparation of those specimens for identification,
can take up to several hours per sample depending on the type of sample, the taxa removed,
the fraction of the material prepared, preparation techniques, and the experience of the
person doing the processing. One worker found that removal of all beetles, Hymenoptera,
and spiders from flight-intercept traps in Montana (without any attempt to separate
families) took from 40 minutes to 5.7 hours per trap depending on the experience of the
individual doing the work (M.A. Ivie, pers. comm.). When studying abundant groups, such as
many families of Diptera, it is unlikely to be practical to prepare all the specimens in a
sample, and subsampling is therefore necessary. A subsample of 8,000 Brachycera (higher
Diptera) from one summers pan-trap samples from an Ontario old-growth forest
required about 800 hours of student assistance to sort, mount and label. Estimates of the
time required for processing and sorting specimens from given trap samples therefore are
very difficult to arrive at, but some minimum estimates are indicated in Table 2. Note the
caveats given in the caption, and also the fact that identification during such processing
would be to the family level only; in most instances, further identification to the
generic or species level would be made by professional systematists.
For any single site, estimated monthly processing time using the recommended sampling
protocol for terrestrial arthropod inventory amounts to 504 hours. Using biology
undergraduate students at a rate of $10 per hour, the cost of processing material
collected at this one site per month would be $5,040. Thus for any single-site inventory
carried out in most areas of Canada, processing for the seven-month period from April to
October would cost $35,280. Compared to the processing costs other expenses (supplies,
travel) are minimal, but note that these processing costs are incurred to reach only the
stage at which species identification becomes possible.
2. Estimates of
minimum time required to process selected samples from one site using recommended sampling
methods and protocol (see Table 3). These minimum
estimates apply only to experienced sorters, sorting specimens from a small number of
selected families (or subsamples when specimens are numerous) to the family level only.
Beginning sorters take much longer, but steadily reduce the time as they gain
experience. Based chiefly on information provided by Scudder (in press).
time required (hrs) per sample
At present, there are very few systematists both willing and able to
identify large numbers of specimens in support of biodiversity studies,
especially because the numbers of professional systematists in Canada
and elsewhere, both in government and universities, and support for
systematic biology in general, have declined greatly in recent years
(Hunter 1991; Wiggins 1992; Heraty 1992). In other words, there is a
shortage of highly trained specialists, and their willingness to
participate in a study should not be taken for granted.
systematists require that specimens submitted for identification be
appropriately prepared and properly labelled, and this takes substantial
time, effort and expertise. Good samples of well prepared material
facilitate accurate identifications and may subsequently prove of value
in the personal research programme of participating systematists.
Adequate numbers of specimens always should be submitted from the
various sampling sites and times of year to permit the recognition of
rare species. For some taxa, submission of subsamples or selected
representatives only should be avoided. Listings of systematists are
available in Arnett and Arnett (1993), Ananthakrishnan (1991) and, for
Coleoptera, Noonan et al. (1993). Other specialized listings are
available. Coddington et al. (1991) have proposed the establishment of a
global network of systematists under the auspices of the IUBS-SCOPE-UNESCO
Program on biodiversity. Listings of arthropod systematists willing to
receive material appear periodically in the Newsletter of the Biological
Survey of Canada (Terrestrial Arthropods). However, the fact that a
systematist exists does not indicate ability or willingness to become
involved in a study. Any prospective participants should be consulted
well in advance of the initiation of sampling and made aware of what is
expected of them as well as of the logistics and ultimate goals of the
survey. Such systematists can provide constructive suggestions as to the
systematists are able to do identifications simply because it is part of
their job, but more often some kind of incentive is required in the form
of coauthorship, specimens of use in the systematist’s research,
reciprocal identifications, or financial compensation for the time and
resources involved. In particular, as emphasized above for the treatment
of unnameable species, specific support for systematics resources should
be included in plans for biodiversity studies, because it will often be
necessary to fund professionals, post-doctoral associates, or students
directly to ensure that identifications will be made, rather than
relying on mainly volunteer effort.
Once target groups have been decided, sampling methods can be chosen.
The second part of this brief provides information relevant to this
choice, and also indicates the sorts of techniques necessary for an
appropriate general inventory.
frame and Follow-up
Results from a given study of biodiversity are useful only if they reach
a stage at which the information can be communicated and used to add to
the store of knowledge about the taxa and sites being studied.
Therefore, a realistic time frame to complete the study, and appropriate
reference materials, must be established.Planning a realistic time frame
is especially important because if resources to complete all aspects of
a biodiversity study are not budgeted, much of the initial effort may be
wasted; it is easier to sample than sort, easier to sort than identify,
and so on. Therefore, the most difficult stages of a project to complete
and to fund are the later ones, comprising identification, publication
of results, and curation of specimens; yet these three elements are
especially critical. Identification to species not only provides names
that allow information to be recorded for future use, but also gives
access to existing information for comparison. Publication of results
forces unfinished analyses to be completed and anecdotal ideas to be
validated, and makes information available in a standardized format that
is much more reliable than some sort of informal report. Deposition and
curation of voucher specimens in museums – which have a long-term
commitment to specimen maintenance – provides reference material for
both ecological information (species associated with particular
habitats, for example) and taxonomic information (Danks et al. 1987).
Future taxonomic study may improve the understanding of particular taxa;
if voucher specimens exist, the specimens can be re-examined to ensure
that the information associated with them is attached to valid species
arthropods: General guidelines
Either active or passive methods can be used to sample arthropods.
Active methods require collection by an individual using various kinds
of equipment. Passive methods establish specialized types of traps at
sampling stations in the field, which are serviced at given intervals.
Passive traps collect large numbers of specimens, and generally remove
the bias introduced by the different abilities of individuals to collect
specimens by active methods. Passive sampling is less labour intensive,
and also captures many species not commonly taken by active collection
(although active collecting is very valuable too in some groups). For
these reasons, passive sampling methods are most frequently used in
sampling arthropod biodiversity.
passive traps left in the field for a period of time require the use of
a preservative, generally a fluid. A solution in water of common salt
and detergent (wetting agent) is most commonly used for frequently
serviced traps, and ethylene or propylene glycol is commonly used for
traps that must be left for long periods or traps in desiccating
environments. Photographic soap is a powerful, non-scented wetting
agent, and automobile antifreeze [“Prestone”] mixed 1:1 with water
provides a satisfactory alternative to expensive laboratory grade
ethylene glycol. Different brands of antifreeze have different
properties, so standardization is important. Unfortunately, ethylene
glycol is highly toxic and attractive to vertebrates, a potential
problem which can be solved by the addition of a bitter substance such
as quinine sulfate or by the use of propylene glycol. If possible, it is
preferable to sample more frequently so that toxic preservatives are not
required. For most temperate environments, salt solutions give adequate
protection from decay for about one week. Obviously, metal containers
should not be used with salt solutions.
passive trapping methods require the removal of specimens from the trap
and their placement in a container in preservative until the samples can
be processed. Most trap types can be serviced with minimal habitat
disturbance by removing the specimens using an aquarium net with a fine
mesh (to ensure that all sizes of specimens are retained), then rinsing
the specimens very gently with water prior to storing the sample in 80%
ethanol. The ethanol should be replaced 1 to 2 days later to avoid
excessive dilution by the water added in the cleansing process. Addition
of 5% acetic acid to the ethanol will prevent specimens from becoming
excessively brittle and will facilitate dissections. We do not recommend
the use of methanol or, especially, formalin as preservatives. “Whirl
Pac”, small “Zip-loc” plastic bags, or plastic “margarine”
containers are ideal for the initial or field phase of specimen storage.
Transfer to glass or polypropylene jars can be made later in the
laboratory, ideally at the time fresh ethanol is being added.
methods for use in a study
The scope of any study is limited by the sampling methods chosen.
Southwood (1978) provides descriptions and an extensive and useful
discussion of most techniques for assessing insect populations. Other
useful overviews of arthropod sampling techniques can be found in Martin
(1977), Disney et al. (1982), Canaday (1987), Steyskal et al. (1986),
Gadagkar et al. (1990), and Górny and Grüm (1993). Some of the most
commonly and widely used sampling methods, their limitations and their
relevance to biodiversity studies are discussed below. The taxa most
frequently collected by each technique are noted, and standardized
sampling protocols are proposed for most techniques.
Recommended methods for conducting an inventory of a good cross section
of the arthropod biodiversity of a site are listed in Table
3. These methods include Malaise, flight-intercept, and pan traps,
and the use of behavioural extractors. All methods are easily
standardized, cost- and labour-effective, and provide a diversity of
high quality material (if serviced as suggested). An accurate picture of
diversity requires multiple-year sampling, because time of occurrence
and abundance of many species differs from year to year (compare Owen
methods: passive collection techniques
A diverse sample of winged insects can be taken using traps which collect flying
insects. Various designs of Malaise traps, which essentially are open-sided tents that
intercept flying insects and direct them to some sort of trap head, are readily available,
many based on Townes (1972) design. All take large numbers of insects, either into
fluid or a dry container. Malaise traps are an efficient way to collect most active,
flying groups such as Tachinidae, aculeate Hymenoptera, and Ichneumonidae and are an
excellent complement to substrate sampling methods. They tend to take large numbers of
non-habitat-associated species (tourists or vagile species breeding in nearby
habitats), and require frequent servicing if some of the specimens (especially large
Diptera) are to be kept in adequate condition for specific identification.
Malaise trap samples vary widely depending on trap design (Disney et al. 1982), mesh size
(Darling and Packer 1988), and colour (Roberts 1972). These factors can be corrected for
by using standard designs available from several commercial sources. However, Malaise trap
efficiency also varies with aspects of installation. Factors such as direction of
prevailing winds, likely flight paths, Malaise head position, and even how tightly the
trap is guyed markedly effect both numerical and taxonomic composition of the catch.
Nonetheless, Malaise traps are arguably the most effective way to trap large numbers of
insect species at a given site, but because they sample only the flying stages
effectively, they should be used together with other trapping techniques in attempts at
Malaise traps in combination with pan traps can take overwhelming numbers of specimens.
Finnamore (1994) estimated that one seasons catch in 39 pan traps plus one Malaise
trap placed in a fen complex near Edmonton, Alberta, totalled approximately 1.5 million
Malaise traps with dry trap heads, usually using dichlorvos or pyrethroids as a killing
agent, must be emptied daily, and the samples either processed immediately or stored in a
freezer. Most extensive Malaise trap surveys use trap heads that funnel the catch into
alcohol or some other preservative. Unfortunately, this renders the Lepidoptera and larger
Diptera difficult to identify, but it is satisfactory for most other taxa. If traps cannot
be serviced frequently, foil can be wrapped around the outside of the Malaise head
container to limit evaporation of the preservative.
We recommend the use of 2 Townes-style Malaise traps (Townes 1972) made of fine black
polyester mosquito netting (sold for use in tent fly screens) per site. Trap dimensions
should be those of Townes (1972). Trap heads should contain 80% ethanol with 5% acetic
acid and should be serviced at weekly intervals.
Many flying insects, especially small or weak flying species, drop down when they
encounter a Malaise trap baffle rather than moving up into the trap head. These species
can be collected by putting pan traps under the centre panel of a Malaise trap. Pan traps
set under a Malaise take a substantially different insect assemblage than do Malaise trap
heads (Marshall 1979; OHara 1988). Alternatively, it is possible to simply stretch
an interception panel, usually made of black mesh, above either a series of pan traps
arranged in a linear fashion, an elongate trough, or even heavy-gauge plastic (preferably
yellow in colour) lining a shallow excavation. This kind of trap, frequently termed a
flight-intercept trap (FIT), is especially efficient in dense forest for sampling small,
flying insects such as small beetles that are otherwise difficult to collect (Peck and
Davies 1980). Although similar traps with a pane of glass as the central panel have been
used (termed window traps: Chapman and Kinghorn 1955), we suggest the use of a porous mesh
as this does not divert or otherwise affect the wind currents on which small, weakly
flying insects may float.
Window or intercept traps may also be used above ground level, and modified intercept
traps have been used for sampling insects in forest canopies (Basset 1988). Spill-proof
pan traps for use as suspended traps in forest insect sampling can be made by cutting an
opening on the broad side of a plastic container such as an antifreeze container (Canaday
1987). Intercept-trap efficiency can be increased significantly by treating the
interception panel with a contact pyrethroid insecticide such as permethrin
[Ambush]. Flooding of the traps can be a problem and should be dealt with, as
for pan traps, through the use of roofs, drainage holes, or frequent (at least once per
We recommend the use of fine black polyester mosquito netting (the same as that suggested
for Malaise trap construction), installed with a row of standard yellow pan traps (see
pan traps) sunk flush with the ground under the centre panel. Suggested
dimensions for a standard panel are 1.25 m (4) high by 1.85 m (6) long. We
suggest the use of a clear plastic roof stretched tent-like over the central panel. The
edges of the roof should not extend below the top of the center panel when viewed from the
side. Because the preservative used in the pans under an intercept trap usually has a
large surface area, a preservative that evaporates slowly is preferred. Two traps should
be installed per site and servicing should be at weekly intervals.
An excellent combination of techniques for long-term survey sites would be a Townes-style
Malaise trap (Townes 1972) made of fine black polyester mosquito netting (sold for use in
tent fly screens), installed with a row of pan traps under the center panel. This combines
the advantages of pan trapping, intercept trapping, and Malaise trapping in a replicable
Pan traps provide a very simple, inexpensive sampling technique using shallow pans of
fluid, usually set into the substrate such that the lip of each pan is flush with the
substrate surface. Pan traps rely on the organism falling or flying into the fluid
preservative; collection may be accidental or the insects may be attracted to the colour
of the pan or, apparently, to the fluid surface. Several commercially available
containers, such as microwave trays, organizer trays, and food distribution containers
make satisfactory pan traps. If studies are to be comparable with one another then pan
shape, surface area, colour, material, edge characteristics, repellent or attractive
properties of trap fluid, and depth of installation must be standardized.
Traps installed flush with the soil surface differ from traps installed flush with the
litter surface (Greenslade 1964), and traps sitting on the substrate surface (sometimes
called water traps) collect a different fauna than those set into the substrate (Disney et
al. 1982). Traps sunk into the substrate take a wider variety of taxa and larger numbers
of specimens. Different taxa are attracted to different trap colours (see for example,
Finch 1991). Yellow seems to be the most widely used colour at the present time and is
especially attractive to various groups of Homoptera and Hymenoptera, although white is
more attractive to some Diptera (Disney et al. 1982). The material from which the trap is
made can affect trap efficiency (Luff 1975); smooth surfaces such as plastic are more
efficient than surfaces that tend to become pitted, such as metal. Toxicity, surface
tension, and other attributes of trap fluid also influence trap efficiency. Because
different taxa are differentially sampled by pan and pitfall traps (Topping 1993), samples
of this sort do not necessarily show a close correlation with density.
Although flooding is not often a problem in frequently serviced traps, the addition of a
roof prevents traps from being flooded by rain. Such flooding will lead to the loss and
degradation of specimens. The addition of roofs must be carefully considered as they add
an extra variable to a sampling programme. Roofs attract some organisms seeking shelter,
and exclude other organisms that may be unwilling to enter a confined area. Drainage holes
provide an alternative method of avoiding flooding. Small, screened, holes near the lip of
a trap generally will prevent fluid overflow. Pan traps can be serviced with minimal
habitat disturbance by removing the specimens using a fine-mesh aquarium net, then rinsing
the specimens very gently with water prior to storing the sample in alcohol.
We recommend rectangular plastic food packaging trays, such as the 500 ml, 15 x 17 cm,
Showcase trays distributed by Edmeads Packaging, Kitchener, Ontario. This type
of readily available product provides a low cost, highly effective pan trap. The outside
of these translucent trays should be spray-painted yellow. Traps should be sunk flush with
the substrate surface. Current inventory programmes use 4-6 such traps per site;
ecological studies will use 10 or more traps, laid out on a grid, per habitat type
sampled. Preservative should be salt solution with soap and servicing should be at weekly
intervals. Little is known about the effect of trap spacing, a factor which probably will
be governed by microhabitat type and variability. Snider and Snider (1986) varied trap
spacing from 0.5-4 m in a northern Michigan forest, but found little effect on catches of
ground beetles or springtails. Nevertheless, it is probably better to keep traps 5 to 10 m
Pan traps collect a large variety of terrestrial arthropods including large numbers of
small arthropods restricted to or living near the substrate surface. Recent pan-trap
surveys of peatlands in Canada have yielded more than 2,000 species of insects in a single
years sampling (Finnamore 1994; Blades and Marshall 1994).
Taxa efficiently sampled using pan traps include spiders, springtails, ground beetles,
sphaerocerid flies, mycetophilid flies, alate aphids, leafhoppers, seed bugs, and several
Although the term pitfall trap is sometimes used to refer to any trap sunk in the ground,
we here restrict the term to traps of relatively small diameter which are deeper than pan
traps and generally placed so that the lip of the trap is flush with the ground surface.
Pitfall traps can be simple containers sunk into the substrate or they can be more
elaborate devices with funnels directing specimens into the trap (Clark 1992; Rivard
1962). Pitfall traps may have fluid preservative in the bottom of the container, or they
may be kept dry or lined with damp paper if living specimens are desired. Traps may be
baited, such as with dung or carrion, but this may render them of limited use for
quantitative studies (Southwood 1978). Baited pitfall traps do, however, offer an
efficient technique for qualitative comparisons of particular communities among sites
(Anderson 1982). Unbaited pitfall traps have been extensively used in sampling
ground-dwelling beetles, particularly carabids (Dennison and Hodkinson 1984; Luff 1975;
Baars 1979). A partial listing of the extensive literature on pitfall traps can be found
in Dunn (1989).
We recommend the use of plastic 450 ml (16 oz.) beer glasses. Four to 10 traps
should be installed per site with covers (15-30 cm (6 to 12")-square pieces of
plywood raised at the corners and weighted with a stone). The preservative should be salt
solution with soap and servicing should be at weekly intervals.
Taxa collected by unbaited traps include many ground-dwelling beetles, spiders, and some
springtails and mites. Taxa collected by baited traps depend on the type of bait used
(Greenslade and Greenslade 1971; Newton and Peck 1975). Some types of baited traps will
attract large vertebrates and precautions must be taken to exclude these scavengers.
Unbaited pitfall traps offer an advantage over simple pan traps in that they permit the
collection of living specimens which can be counted, identified and released or used in
Several authors have argued that light traps are the best way to undertake insect surveys
(Holloway 1980; Gadagkar et al. 1990), because they collect well known taxa efficiently,
they collect large numbers of insects, and they have been in wide use for a longer period
than other mass-trapping devices. Light traps certainly are one of the most powerful
collecting tools available, but a large number of variables affects the size and taxonomic
composition of light-trap catches (Bowden 1982). Trap size, height, design, surroundings,
and bulb type are some of the variables that should be considered. Diptera, for example,
are taken in larger numbers in incandescent traps and Lepidoptera are more abundant in
ultra-violet (UV) traps (Southwood 1978). Mercury-vapour light traps will attract a
somewhat different fauna than ultra-violet lights. Most light traps use a killing agent
such as cyanide, Vapona or ethyl acetate; however, an electrified trap designed by
Mizutani et al. (1982) bypasses the need for highly toxic killing agents. Some light-trap
designs collect insects directly into a fluid preservative, which is suitable for groups
such as Hymenoptera, Coleoptera and Psocoptera but undesirable for Lepidoptera and
Diptera. A combination of electricity and a safe insecticide, such as a pyrethroid, might
be the best combination. Because of its use in one of the most important insect surveys
done to date (the Rothamsted Insect Survey: Taylor and French 1974) the Rothamsted light
trap (Williams 1948) is recommended as an appropriate design for arthropod biodiversity
Light traps, although widely used, collect only certain, vagile taxa (many of which may be
tourists) which are active at night. Often they require nightly setup and
servicing. Taxa collected include primarily Lepidoptera, Coleoptera, some Diptera,
Neuroptera, Hymenoptera, and Hemiptera.
Emergence Traps and Tent Traps
Traps that surround or cover a unit of habitat, and subsequently collect arthropods that
move upwards towards the light, can collect a wide variety of insects directly associated
with a specific habitat unit. While most studies using this kind of trap have dealt with
specific taxa such as pest leafhoppers (Cherry et al. 1977), such devices are of potential
value for biodiversity studies. Rosenberg et al. (1987) used a modified LeSage and
Harrison aquatic emergence trap (LeSage and Harrison 1979) to sample arthropods in an
Ontario peatland. This proved efficient for the collection of Chironomidae, but, for
instance, took only 12 species of Sphaeroceridae. Pan traps in similar peatlands would be
expected to take 40-50 species of Sphaeroceridae (Marshall 1994). Emergence or tent traps
collect only positively phototactic arthropods present on or in the habitat at the time of
installation of the trap, which can be an advantage or disadvantage depending on the goals
of the project. These methods collect only resident species and can take large numbers of
specimens. Adis and Schubart (1985) used tent traps (ground photo-eclectors)
to collect between 1,000 and 7,000 specimens (mostly Diptera) per square metre in a
central Amazonian forest, compared to between 30 and 60 specimens per square metre
collected in the forest canopy.
We recommend these types of traps for specialized studies where the fauna emerging from a
particular type of habitat or vegetation is being analyzed. Emergence traps can be placed
over a portion of a tree, for instance, to survey bark beetles emerging from the tree
trunk or limbs.
Traps that take insects which come into contact with an adhesive surface are widely used
for sampling pest taxa such as stable flies (Williams 1973), mosquitoes (Dow and Morris
1972), and aphids (ABrook 1973). Their efficiency is affected by colour, size,
orientation, shape, environmental conditions, and type of adhesive. Grease-based adhesives
are more easily dissolved than resin-based adhesives, but only weak insects are trapped
effectively by grease (Southwood 1978). Even with the easily dissolved grease-based
adhesives, specimens are difficult to handle and the quality of the specimens collected
often is well below the level required for specific identification of many taxa. We feel
that flight-intercept traps are better alternatives to sticky traps when identification of
a variety of taxa is an important component of a project. We do not recommend the general
use of sticky traps in biodiversity studies.
Sampling methods: active collection
Extraction of invertebrates from substrate samples can yield large numbers of wingless
arthropods. Of special interest and value is the frequent capture and thus association of
conspecific adults and immature stages. Despite this attractive feature, identification of
immature stages to the species level is difficult and in many instances it is not
possible. Substrate sampling is mainly of importance for surveying wingless taxa such as
Collembola, mites, some groups of spiders and some ground-substrate-dwelling groups of
beetles which generally will not be collected through other means. Substrate sampling is
an excellent complement to Malaise/flight intercept/pan trapping. It should be noted that
substrate sampling effectively samples resident species and very few tourists
are represented. The main methods of substrate sampling are collecting by hand,
behavioural extractors, and soil washing and flotation (see Górny and Grün 1993).
Behavioural extractors and soil washing and flotation require active field collection of
samples followed by passive processing to separate the arthropods from the substrate.
Collecting by Hand
The simplest method for sampling the biodiversity of large litter and soil arthropods such
as millipedes, centipedes and some larger active beetles, is to mark a quadrat of 0.5-1.0
m2 (or some other area) in the field and remove the litter and soil progressively, usually
to a depth of 10 cm. The litter and soil can be processed onto a sheet or board through a
range of sifters which are used to break up the debris and separate the arthropods from
it. The arthropods present are simply collected as they move. Organisms which are not
active, even if large, may be incompletely sampled using this method. This process depends
on specimen movement and visual observation by the collector of that movement and results
may be highly subjective and dependent on the observer. An alternative would be to employ
field quadrats to collect the sample but use a behavioural extractor to remove the
Although offering a reliable means of standardization by sampling a defined area of
substrate, the method is labour-intensive and experience shows that randomly selected
quadrats yield low numbers of specimens and low species diversity per unit of sampling
effort regardless of how specimens are extracted. We do not recommend its use for general
biodiversity surveys but we acknowledge that it may be useful for confirmation of
microhabitat use of various taxa.
Behavioural Extractors (Berlese and Tullgren Funnels)
Berlese and Tullgren extractors are methods of separating arthropods from soil and litter,
which generally involve using heat and desiccation to stimulate the animals to leave the
samples on their own (Martin 1977). As a result only active, free-living stages are
extracted. These extractors and their many modifications are the most practical and widely
used methods of assessing the diversity and abundance of smaller, less mobile, cryptic
arthropods in soil and litter. They can also be used successfully to collect arthropods
from loose bark, rotting wood, bracket fungi, mosses, flowers, manure and nests (Martin
1977). The sample unit can be a core or cube, ranging from 2.5 cm to 1 metre in diameter,
or can be a specific volume of substrate (Wright and Coleman 1988). In a survey of 20
recent publications on soil arthropods the most common core size used was 5 cm in
diameter, to a depth of 15 cm in soil. However, the core size used will depend on the size
of arthropods being collected and the habitat (Edwards 1991). In peatland soils, for
example, Borcards (1991) sample size was 15 cm x 15 cm to a depth of 10 cm. Samples
may be collected randomly, or they may be collected using field quadrats.
For inventory purposes a quantifiable volume of substrate is not essential and the size of
the sample unit may depend on the type of microhabitat being assessed. In these instances,
the microhabitat can be concentrated by passing the litter through a sifter (Norton and
Kethley 1988). As these authors note, this eliminates bulky, larger pieces of substrate
and enhances uniform drying of the siftings.
The essential components of these extractors (see Martin 1977; Steyskal et al. 1986) are a
sample container with wire mesh or screening on the bottom, a metal or plastic funnel in
which, or over which, the sample container is placed and a collecting vessel below the
funnel which usually contains a liquid preservative, generally 70-80% ethanol with 5%
acetic acid. A source of heat and desiccation (light bulb, electric resistance wire, or if
necessary, sunlight) is placed above the sample. The objective is to create a steep
gradient of temperature and moisture throughout the sample (Edwards 1991). The arthropods
react to the heat and desiccation by moving downward (away from the heat) and eventually
fall through the screen at the bottom into the preservative (Martin 1977). Normally, soil
cores should not be deeper than 5 cm and should be inverted when placed in the sample
containers. Chemicals (e.g. napthalene) may be used in place of the heat source, but
generally these are not very effective. Cheesecloth below the sample and/or a baffle in
the funnel can be used to reduce debris falling out of the sample as it dries or is
agitated by the movement of larger organisms. The wattage of the light bulb used depends
on the size of the funnel (Martin 1977). The length of time of extraction varies from 6
hours to 1-2 weeks and depends on the moisture content of the sample, the intensity of the
heat source, the depth and uniformity of the sample and the types of arthropods desired.
Mites, for example, and especially immatures, may take a relatively long period of time
(up to 14 days) to exit the sample. When funnels must be operated outdoors a tight-fitting
hood is recommended to avoid contamination of samples by insects attracted to the light
(Martin 1977). Although such species can often be identified as contaminants because their
natural history is known, contamination can also result from specimens hanging on to, or
becoming entangled in, the cheesecloth. To avoid this possibility, the cheesecloth and
wire mesh should be removed after the processing of each sample and such specimens
Commonly used modifications of the Berlese-Tullgren funnel include the Macfadyen
high-gradient funnel, the Kempson apparatus (Kempson et al. 1963), and the
Merchant-Crossley extractor (Merchant and Crossley 1970; Norton 1986). Norton and Kethley
(1988) describe a light-weight, collapsible and easily transportable Berlese funnel made
of rip-stop nylon, which is both simple to construct and efficient. Another design is
presented by Wheeler and McHugh (1987). Edwards (1991) reviews most modifications of
behavioural extractors and provides details of preferred methods for sampling different
We recommend the use of a sifter to concentrate the substrate. Five-litre samples of
sifted litter, up to 10 per site per sampling period, should be collected. In inventory
studies, a uniform effort should be made to sift in various microhabitats (e.g. under
fallen logs, under fungi, under fruit- or seed-fall, etc.). For litter layer arthropods,
such as beetles, a defined area of litter can be sifted for a defined time period.
For a given site, samples should be taken 4 times per year to ensure that active stages of
species quiescent during some parts of the year are represented. All storage and transport
of samples should be in containers which do not permit the buildup of excessively high
temperatures or high humidity. We suggest that each sample be placed in a lightweight
cotton or ripstop nylon bag; pillow cases are ideal.
We recommend that funnels be constructed of lightweight sheet metal or aluminum flashing.
Funnels should be 50 cm (20") high and 35 cm (14") in diameter at the top and
tapered to 2.5-5 cm (1-2") in diameter at the bottom. Hardware cloth of 1.25 cm
(1/2") mesh should be placed in the funnel 15 cm (6") from the top and covered
with single-ply cheesecloth. Samples should be processed for between 8 hours and 14 days
using 60W bulbs suspended 10 cm (4") above the top of the sample.
Soil Washing and Flotation
Behavioural extraction methods extract only the active stages of arthropods, and in arid
soils, deep soils, and mineral soils with high clay content are inefficient for certain
groups, such as endeostigmatic mites and podurid and onychiurid Collembola (Walter et al.
1987). For any inventory in these habitats, and for these groups, soil washing and
flotation (Kethley 1991), or direct heptane flotation of soil cores (Walter et al. 1987)
is recommended. Soil washing often is a difficult technique to use because it requires
large quantities of water (about a 4:1 ratio of water to soil), but as Kethley (1991)
notes this technique provides complete life-history data for many microarthropods and is
the only effective way to assess diversity in deep soils where arthropod numbers are very
low. The recent development of heptane flotation (Walter et al. 1987; Kethley 1991) is
based on the affinity of the arthropod cuticle for petroleum derivatives such as heptane.
This method is useful in some circumstances as it allows large numbers of soil samples to
be stored for some time and processed when convenient. In contrast, processing using
behavioural extractors should be carried out as soon as possible after field collection of
the samples (Edwards 1991).
Projects that attempt to approximate complete inventories of a given habitat or locality,
or projects that require an estimate of the degree to which a trapping programme is
sampling the total fauna, require techniques that collect virtually all the arthropods in
a given unit of habitat. Suction devices such as the widely used D-vac
(Dietrick 1961) are often considered to obtain total faunal samples, but differ in
efficiency for different taxa and for different substrates. Theoretically, a very high
power suction sampler such as the motor-vehicle-sized McCoy Insect Collector
(McCoy and Lloyd 1975) can take almost 100% of the fauna, unfortunately along with a great
deal of substrate. Vacuum samples which include litter, leaves, and other debris must be
laboriously sorted by hand unless some kind of behavioural extractor is used. Behavioural
extraction necessitates that the specimens be kept alive, and even so only a sample of the
arthropods in the collection would be extracted. Vacuum sampling might take some species
missed by passive trapping programmes, but should not be considered as a means of
measuring absolute species richness.
Another approach to total inventory is to make collections by using chemicals to kill or
stun everything in a unit area of habitat. This method has been used to sample fruit-tree
insects (Collyer 1951) and is now used for biodiversity studies in the tropical forest
canopy. The recent (and controversial) estimates of total global biodiversity (Erwin 1982;
Stork 1988) are based largely on chemical knockdown of insects from tropical trees. The
target area is sprayed or fogged, usually with a pyrethroid insecticide, and the affected
organisms fall onto sheets, trays, or funnels placed below. A method for canopy spraying
is described by Martin (1966), the merits of canopy fogging are discussed by Paarmann and
Stork (1987) and Adis et al. (1984) and a summary of the protocol and methodology
underlying insecticide sampling in trees is given by Stork (1988).
Knockdown methods are selective for larger insects that do not stick to the foliage, and
can undercollect smaller insects by as much as 50% (Muir and Gambrill 1960). Much of the
fauna living under bark or in leaves is not sampled. Despite these concerns, this
technique remains the most popular and effective approach to the study of canopy arthropod
biodiversity. Chemical methods do have the advantage of being relatively independent of
insect activity and climatic conditions, and can be applied to specific microhabitats such
as individual trees, specific parts of trees, or a specific volume of canopy. Canopy
fogging in a tropical forest yields thousands of specimens in a matter of a few hours.
Stork (1988) reports 24,000 individuals representing over 2,800 species from only 10 trees
in Borneo. In order to take seasonality into account fogging samples should be taken at
various intervals throughout the year. Immature stages are rarely collected using this
technique unless they feed externally on foliage.
Canopy fogging was originally developed for study of temperate forests; however, it is
most effective in tropical forests where the forest canopy is complex. Most foliage in a
tropical forest is in the canopy and there is high plant-species diversity and an
abundance of epiphytes. This technique is somewhat expensive and complex to set up and to
use effectively. It would be useful for studying the insects of the forest canopy in
particular, but we do not recommend the use of this method in most studies of
biodiversity, particularly at temperate latitudes.
Suction and Rotary Traps
In contrast to Malaise traps, which passively sample flying insects, suction and rotary
traps actively sample the insects in a given volume of air. They do this either by pumping
a volume of air through a filter (Johnson 1950) or by using a mechanically rotated net
that continuously samples aerial fauna (Chamberlin 1940). Taylor (1962) suggests that
these traps sample 85% of the flying population. Suction traps collect slowly and weakly
flying insects which form a kind of aerial plankton. Certain elements of the fauna are
easily collected in this manner. Most studies using these types of traps deal only with
specific pest species (Taylor 1962), or list general collections identified only to order
We consider that suction traps constitute an effective alternative method for selected
faunal inventory projects. Suction traps collect a number of small, fragile, winged
insects which are otherwise not sampled. Among the groups taken in suction traps are
micro-Coleoptera, micro-Hymenoptera, Coniopterygidae (Neuroptera), and alate aphids
(Homoptera). One drawback of both suction and rotary traps is that an electrical power
outlet must be near the habitat. We do not recommend the general use of rotary traps.
Sweeping and Beating Vegetation
The use of an insect net is the most commonly and widely known technique for collecting
insects. As a routine sampling method, the use of a net is appropriate in some habitats,
but only under uniform conditions. Weather, vegetation type and age, weight of net, type
of mesh, and handler skill are some of the factors affecting net collections. Those
relatively few taxa which sit high on the vegetation and do not fall off when approached
might be efficiently sampled, and some of them may include species rarely collected in
Malaise or pan traps. A measure of numbers of sweeps can be used for standardization, but
because of high user bias this method is recommended only as an auxiliary sampling
technique in an inventory project. If sweep netting is used we suggest that the standard
dimensions of the net be 38 cm and that 20 sweeps of 180° constitute a sample.
Another commonly used method of general collecting is to place a sheet under a plant and
to strike the vegetation so that the arthropods on the plant are dislodged and fall onto
the sheet. Such beating sheets generally are supported by a framework of interlocking
lightweight poles and are held under the plant with one hand while the plant is struck
with a stick held in the other hand. As with the use of nets, standardization is possible
but subject to bias; beating sheets are recommended only as an auxiliary sampling
technique in an inventory project. If beating is used we suggest that the standard
dimensions of the sheet be 1 m2 and that 20 beats per tree or sample be made.
Specialists in every group of arthropods prefer particular, often specialized, methods of
collecting. Specialized techniques can be used to supplement baseline inventory data
gathered using the sampling methods discussed above, or they can be used for biodiversity
measures using limited collections of particular taxa. Coddington et al. (1992) argue that
hit and run sampling trips, or single-visit collecting trips, are the only
practical way to sample tropical biodiversity. Given this premise, they offer an
assessment of looking up, looking down, beating, and sifting as
quantifiable methods of sampling spiders. This kind of spot assessment of selected taxa
using selected techniques will probably remain the only practical approach to biodiversity
assessment for much of the tropics, and offers many efficiencies over exhaustive
inventories. For example, dealing with large numbers of common species is a major cost of
processing mass samples. Specialized collecting usually allows for the rejection of common
species after a certain number have been collected, thus lowering the cost of processing
the resultant collections.
The main role of specialized collecting in broader biodiversity studies is to supplement,
and assess the efficiency of, mass sampling devices such as pan traps and behavioural
extractors. Specialists on phytophagous taxa are almost certain to find additional species
by beating specific plants, searching appropriate hosts, and by rearing from hosts or
plant parts. Similarly, specialists are able to recognise microhabitats from which they
can net, aspirate or otherwise hand collect rare species which might be missed by other
techniques. Members of the acarine suborder Oribatida, for example, are commonly
associated with soil and litter, but any inventory of this group should include
specialized collecting such as twig washing, leaf washing, observation of leaf surfaces,
and collections from the axils of twigs and branches. It is therefore important for any
inventory project to allow for site visits by specialists in the taxa under consideration.
Such visits can generate additional taxon records, and cast light on trapping efficiency
and habitat heterogeneity from different perspectives.
The most time-consuming aspect of any typical biological
inventory or biodiversity study is the conversion of raw samples into
prepared, labelled and sorted lots to be identified in house or sent off for specific
identification. It is essential that instructions for how to prepare specimens be
solicited from cooperating systematists and that these be strictly adhered to. Specimens
trapped in fluid usually must be dried in a critical-point drier (Gordh and Hall 1979),
carefully mounted to the specifications of the cooperating systematist, labelled using
appropriate data and paper (Darling and Plowright 1990), then sorted to sendable
units, usually family, and directed to the appropriate cooperating systematist. In
some cases, material will have to be mounted on slides or sorted into alcohol. The
importance of proper preparation cannot be overemphasized. For example, specific
identification of 500 properly point-mounted, critical-point-dried flies might take a
specialist about one week, a good investment of time if the data on the specimens rendered
them useful. If the specimens were air dried, identification time might increase to four
weeks and might render the investment of time impractical. If the submitted flies were not
glued to points firmly enough to allow dissection without remounting, it could again
double the time required for identification.
Such specialized demands must be taken into
account when preparing a budget for any inventory project. One hour a week spent emptying
pan traps can easily keep a full-time technician busy preparing and sorting material. The
ratio between sampling costs and processing costs can be as high as 1:40.
A well designed project is likely to result in the collection
of tens of thousands of specimens, some of which will become valuable identified voucher
specimens, and some of which will be unidentifiable at the present time, yet worthy of
retention. It is important to consider the long term integrity of this material. As
emphasized for the initial planning of a study (above), voucher specimens must be placed
in museums with a long-term commitment to specimen maintenance. Unidentified material
should be maintained with appropriate information for later incorporation into, or
comparison with, the project database. Consideration of the cost of storage materials such
as pins, unit trays, drawers, insect cabinets, and subsequent curatorial activity
therefore should be an integral part of any inventory budget.
It is important to establish appropriate databases to allow
for maintenance and exchange of data files on habitats and specimens. Computerization of
data, including unique specimen codes, can greatly facilitate database usage and future
additions to the database. There is a need for systems that allow specimens to be readily
tracked and related back to a particular site and date. Janzen (1991) has pioneered the
use of bar-coded specimen labels for this purpose. The use of bar codes can be costly and
may be best suited for inventories in which species diversity is high and the number of
individuals of many of the species collected is low. Standard use of precise latitude and
longitude data for sample sites, notably by using Geographical Positioning System (GPS)
capabilities, is highly desirable.
An extensive and developing literature considers requirements for specimen databases. For
example, standards for fields and terms for collection data for insects have been proposed
by Noonan (1990) and Noonan and Thayer (1990). These aspects of reporting and tracking
information on specimens collected during biodiversity inventories are important for the
long-term value of any study, but details of such requirements are beyond the scope of
It has been estimated that at
present only about half of the insect fauna of Canada is known. Regional inventories, done
properly and supported by an adequate systematics infrastructure, can work towards
resolving this lack of knowledge while at the same time providing the baseline data needed
for proper and efficient management of biodiversity. Such inventories are expensive and
time consuming to carry out, because the numbers of arthropod species and specimens
collected greatly exceed the numbers resulting from surveys of other taxa.
Nevertheless, surveys of arthropod biodiversity are feasible, given proper planning and
suitable sampling protocols, as explained in this brief. We re-emphasize that the limiting
factor in any inventory of arthropod diversity is the strength of the systematics
resources available to support, and in many cases carry out, this kind of work.
Consequently, much of the biodiversity in our own backyard will remain unknown unless the
numbers of systematists and the support for this science increase.
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This brief was prepared by a subcommittee of
the Biological Survey of Canada:
S.A. Marshall, R.S. Anderson, R.E. Roughley, V.
Behan-Pellletier, and H.V. Danks.
Originally published by the Entomological Society of
Supplement to the Bulletin, Vol. 26 (1), March 1994