The Insects - Outline of Entomology 3th Edition - Chapter 17 docx

Active collecting involves search-ing the environment for insects, and may be preceded by periods of observation before obtaining specimens for identification purposes.. Such techniques

METHODS IN ENTOMOLOGY:

COLLECTING, PRESERVATION, CURATION, AND IDENTIfiCATION

Alfred Russel Wallace collecting butterflies (After various sources, especially van Oosterzee 1997; Gardiner 1998.)

For many entomologists, questions of how and what to

collect and preserve are determined by the research

project (see also section 13.4) Choice of methods may

depend upon the target taxa, life-history stage,

geo-graphical scope, kind of host plant or animal, disease

vector status, and most importantly, sampling design

and cost-effectiveness One factor common to all such

studies is the need to communicate the information

unambiguously to others, not least concerning the

identity of the study organism(s) Undoubtedly, this

will involve identification of specimens to provide

names (section 1.4), which are necessary not only to

tell others about the work, but also to provide access to

previously published studies on the same, or related,

insects Identification requires material to be

appropri-ately preserved so as to allow recognition of

morpho-logical features which vary among taxa and life-history

stages After identifications have been made, the

speci-mens remain important, and even have added value,

and it is important to preserve some material (

vouch-ers) for future reference As information grows, it may

be necessary to revisit the specimens to confirm

iden-tity, or to compare with later-collected material

In this chapter we review a range of collecting

methods, mounting and preservation techniques, and

specimen curation, and discuss methods and principles

of identification

17.1 COLLECTION

Those who study many aspects of vertebrate and plant

biology can observe and manipulate their study

organ-isms in the field, identify them and, for larger animals,

also capture, mark, and release them with few or no

harmful effects Amongst the insects, these techniques

are available perhaps only for butterflies and

dragon-flies, and the larger beetles and bugs Most insects can

be identified reliably only after collection and

preserva-tion Naturally, this raises ethical considerations, and it

is important to:

• collect responsibly;

• obtain the appropriatepermit(s);

• ensure that voucher specimensare deposited in a

well-maintained museum collection

Responsible collecting means collecting only what is

needed, avoidance or minimization of habitat

destruc-tion, and making the specimens as useful as possible to

all researchers by providing labels with detailed

collec-tion data In many countries or in designated reserve

areas, permission is needed to collect insects It is the collector’s responsibility to apply for permits and fulfill the demands of any permit-issuing agency Further-more, if specimens are worth collecting in the first place, they should be preserved as a record of what has been studied Collectors should ensure that all speci-mens (in the case of taxonomic work) or at least repres-entative voucher specimens (in the case of ecological, genetic, or behavioral research) are deposited in a recognized museum Voucher specimens from surveys

or experimental studies may be vital to later research Depending upon the project, collection methods may

be active or passive Active collecting involves search-ing the environment for insects, and may be preceded

by periods of observation before obtaining specimens for identification purposes Active collecting tends to

be quite specific, allowing targeting of the insects to

be collected Passive collecting involves erection or installation of traps, lures, or extraction devices, and entrapment depends upon the activity of the insects themselves This is a much more general type of collect-ing, being relatively unselective in what is captured

17.1.1 Active collecting

Active collecting may involve physically picking indi-viduals from the habitat, using a wet finger, fine-hair brush, forceps, or an aspirator(also known in Britain

as a pooter) Such techniques are useful for relatively slow-moving insects, such as immature stages and sedentary adults that may be incapable of flying or reluctant to fly Insects revealed by searching particu-lar habitats, as in turning over stones, removing tree bark, or observed at rest by night, are all amenable

to direct picking in this manner Night-flying insects can be selectively picked from a light sheet– a piece

of white cloth with an ultraviolet light suspended above

it (but be careful to protect eyes and skin from exposure

to ultraviolet light)

Netting has long been a popular technique for capturing active insects The vignette for this chapter depicts the naturalist and biogeographer Alfred Russel

Wallace attempting to net the rare butterfly, Graphium androcles, in Ternate in 1858 Most insect nets have a

handle about 50 cm long and a bag held open by a hoop

of 35 cm diameter For fast-flying, mobile insects such

as butterflies and flies, a net with a longer handle and

a wider mouth is appropriate, whereas a net with a narrower mouth and a shorter handle is sufficient for

small and/or less agile insects The net bag should

always be deeper than the diameter so that the insects

caught may be trapped in the bag when the net is

twisted over Nets can be used to capture insects whilst

on the wing, or by using sweeping movements over the

substrate to capture insects as they take wing on being

disturbed, as for example from flower heads or other

vegetation Techniques of beating (sweeping) the

vegetation require a stouter net than those used to

intercept flight Some insects when disturbed drop to

the ground: this is especially true of beetles The

tech-nique of beating vegetation whilst a net or tray is held

beneath allows the capture of insects with this

defen-sive behavior Indeed, it is recommended that even

when seeking to pick individuals from exposed

posi-tions, that a net or tray be placed beneath for the

inevitable specimen that will evade capture by

drop-ping to the ground (where it may be impossible to

locate) Nets should be emptied frequently to prevent

damage to the more fragile contents by more massive

objects Emptying depends upon the methods to be used

for preservation Selected individuals can be removed

by picking or aspiration, or the complete contents can

be emptied into a container, or onto a white tray from

which targeted taxa can be removed (but beware of fast

fliers departing)

The above netting techniques can be used in aquatic

habitats, though specialist nets tend to be of different

materials from those used for terrestrial insects, and of

smaller size (resistance to dragging a net through water

is much greater than through air) Choice of mesh size

is an important consideration – the finer mesh net

required to capture a small aquatic larva compared

with an adult beetle provides more resistance to being

dragged through the water Aquatic nets are usually

shallower and triangular in shape, rather than the

cir-cular shape used for trapping active aerial insects This

allows for more effective use in aquatic environments

17.1.2 Passive collecting

Many insects live in microhabitats from which they are

difficult to extract – notably in leaf litter and similar soil

debris or in deep tussocks of vegetation Physical

inspection of the habitat may be difficult and in such

cases the behavior of the insects can be used to separate

them from the vegetation, detritus, or soil Particularly

useful are negative phototaxic and thermotaxic and

positive hygrotaxic responses in which the target

insects move away from a source of strong heat and/or light along a gradient of increasing moisture, at the end

of which they are concentrated and trapped The

Tullgren funnel(sometimes called a Berlese funnel) comprises a large (e.g 60 cm diameter) metal funnel tapering to a replaceable collecting jar Inside the funnel a metal mesh supports the sample of leaf litter

or vegetation A well-fitting lid containing illuminating lights is placed just above the sample and sets up a heat and humidity gradient that drives the live animals downwards in the funnel until they drop into the collecting jar, which contains ethanol or other preservative

The Winkler bag operates on similar principles, with drying of organic matter (litter, soil, leaves) forcing mobile animals downwards into a collecting chamber The device consists of a wire frame enclosed with cloth that is tied at the top to ensure that speci-mens do not escape and to prevent invasion by scav-engers, such as ants Pre-sieved organic matter is placed into one or more mesh sleeves, which are hung from the metal frame within the bag The bottom of the bag tapers into a screw-on plastic collecting jar con-taining either preserving fluid or moist tissue paper for live material Winckler bags are hung from a branch or from rope tied between two objects, and operate via the drying effects of the sun and wind However, even mild windy conditions cause much detritus to fall into the residue, thus defeating the major purpose of the trap They are extremely light, require no electric power and are very useful for collecting in remote areas, although when housed inside buildings or in areas subject to rain

or high humidity, they can take many days to dry com-pletely and thus extraction of the fauna may be slow

Separating bags rely on the positive phototaxic (light) response of many flying insects The bags are made from thick calico with the upper end fastened to

a supporting internal ring on top of which is a clear Perspex lid; they are either suspended on strings or supported on a tripod Collections made by sweeping

or specialized collections of habitat are introduced by quickly tipping the net contents into the separator and closing the lid Those mobile (flying) insects that are attracted to light will fly to the upper, clear surface, from which they can be collected with a long-tubed aspirator inserted through a slit in the side of the bag Insect flight activity is seldom random, and it is pos-sible for the observer to recognize more frequently used routes and to place barrier traps to intercept the flight path Margins of habitats (ecotones), stream lines, and

gaps in vegetation are evidently more utilized routes.

Traps that rely on the interception of flight activity and

the subsequent predictable response of certain insects

include Malaise traps and window traps The Malaise

trapis a kind of modified tent in which insects are

inter-cepted by a transverse barrier of net material Those

that seek to fly or climb over the vertical face of the trap

are directed by this innate response into an uppermost

corner and from there into a collection jar, usually

containing liquid preservative A modified Malaise

trap, with a fluid-filled gutter added below, can be used

to trap and preserve all those insects whose natural

reaction is to drop when contact is made with a barrier

Based on similar principles, the window trapconsists

of a window-like vertical surface of glass, Perspex, or

black fabric mesh, with a gutter of preserving fluid lying

beneath Only insects that drop on contact with the

window are collected when they fall into the preserving

fluid Both traps are conventionally placed with the

base to the ground, but either trap can be raised above

the ground, for example into a forest canopy, and still

function appropriately

On the ground, interception of crawling insects can

be achieved by sinking containers into the ground to

rim-level such that active insects fall in and cannot

climb out These pitfall traps vary in size, and may

feature a roof to restrict dilution with rain and preclude

access by inquisitive vertebrates (Fig 17.1) Trapping

can be enhanced by construction of a fence-line to

guide insects to the pitfall, and by baiting the trap

Specimens can be collected dry if the container

con-tains insecticide and crumpled paper, but more usually

they are collected into a low-volatile liquid, such as pro-pylene glycol or ethylene glycol, and water, of varying composition depending on the frequency of visitation

to empty the contents Adding a few drops of detergent

to the pitfall trap fluid reduces the surface tension and prevents the insects from floating on the surface of the liquid Pitfall traps are used routinely to estimate species richness and relative abundances of ground active insects However, it is too rarely understood that strong biases in trapping success may arise between compared sites of differing habitat structure (density

of vegetation) This is because the probability of capture

of an individual insect (trappability) is affected by the complexity of the vegetation and/or substrate that sur-rounds each trap Habitat structure should be meas-ured and controlled for in such comparative studies Trappability is affected also by the activity levels of insects (due to their physiological state, weather, etc.), their behavior (e.g some species avoid traps or escape from them), and by trap size (e.g small traps may

exclude larger species) Thus, the capture rate (C) for pitfall traps varies with the population density (N) and trappability (T ) of the insect according to the equation

C = TN Usually, researchers are interested in

estimat-ing the population density of captured insects or in determining the presence or absence of species, but such studies will be biased if trappability changes between study sites or over the time interval of the study Similarly, comparisons of the abundances of different species will be biased if one species is more trappable than another

Many insects are attracted by baitsor lures, placed

in or around traps; these can be designed as “generic”

to lure many insects, or “specific”, designed for a single target Pitfall traps, which trap a broad spectrum of mobile ground insects, can have their effectiveness increased by baiting with meat (for carrion attraction), dung (for coprophagous insects such as dung beetles), fresh or rotting fruit (for certain Lepidoptera, Coleop-tera, and Diptera), or pheromones (for specific target insects such as fruit flies) A sweet, fermenting mixture

of alcohol plus brown sugar or molasses can be daubed

on surfaces to lure night-flying insects, a method termed “sugaring” Carbon dioxide and volatiles such

as butanol can be used to lure vertebrate-host-seeking insects such as mosquitoes and horseflies

Colors differentially attract insects: yellow is a strong lure for many hymenopterans and dipterans This behavior is exploited in yellow pan traps which are simple yellow dishes filled with water and a

surface-Fig 17.1 A diagrammatic pitfall trap cut away to show

the inground cup filled with preserving fluid (After an

unpublished drawing by A Hastings.)

tension reducing detergent and placed on the ground to

lure flying insects to death by drowning Outdoor

swimming pools act as giant pan traps

Light trapping(see section 17.1.1 for light sheets)

exploits the attraction to light of many nocturnal flying

insects, particularly to the ultraviolet light emitted by

fluorescent and mercury vapor lamps After attraction

to the light, insects may be picked or aspirated

indi-vidually from a white sheet hung behind the light, or

they may be funneled into a container such as a tank

filled with egg carton packaging There is rarely a need

to kill all insects arriving at a light trap, and live insects

may be sorted and inspected for retention or release

In flowing water, strategic placement of a stationary

net to intercept the flow will trap many organisms,

including live immature stages of insects that may

otherwise be difficult to obtain Generally, a fine mesh

net is used, secured to a stable structure such as bank,

tree, or bridge, to intercept the flow in such a way that

drifting insects (either deliberately or by dislodgement)

enter the net Other passive trapping techniques in

water include emergence traps, which are generally

large inverted cones, into which adult insects fly on

emergence Such traps also can be used in terrestrial

situations, such as over detritus or dung, etc

17.2 PRESERVATION AND CURATION

Most adult insects are pinned or mounted and stored

dry, although the adults of some orders and all

soft-bodied immature insects (eggs, larvae, nymphs, pupae

or puparia) are preserved in vials of 70 – 80% ethanol

(ethyl alcohol) or mounted onto microscope slides

Pupal cases, cocoons, waxy coverings, and exuviae

may be kept dry and either pinned, mounted on cards

or points, or, if delicate, stored in gelatin capsules or in

preserving fluid

17.2.1 Dry preservation

Killing and handling prior to dry mounting

Insects that are intended to be pinned and stored dry

are best killed either in a killing bottleor tube

con-taining a volatile poison, or in a freezer Freezing avoids

the use of chemical killing agents but it is important to

place the insects into a small, airtight container to

pre-vent drying out and to freeze them for at least 12–24 h

Frozen insects must be handled carefully and properly

thawed before being pinned, otherwise the brittle appendages may break off The safest and most readily available liquid killing agent is ethyl acetate, which although flammable, is not especially dangerous unless directly inhaled It should not be used in an enclosed room More poisonous substances, such as cyanide and chloroform, should be avoided by all except the most experienced entomologists Ethyl acetate killing con-tainers are made by pouring a thick mixture of plaster

of Paris and water into the bottom of a tube or wide-mouthed bottle or jar to a depth of 15 –20 mm; the plaster must be completely dried before use To “charge”

a killing bottle, a small amount of ethyl acetate is poured onto and absorbed by the plaster, which can then be covered with tissue or cellulose wadding With frequent use, particularly in hot weather, the container will need to be recharged regularly by adding more ethyl acetate Crumpled tissue placed in the container will prevent insects from contacting and damaging each other Killing bottles should be kept clean and dry, and insects should be removed as soon as they die to avoid color loss Moths and butterflies should be killed separately to avoid them contaminating other insects with their scales For details of the use of other killing agents, refer to either Martin (1977) or Upton (1991) under Further reading

Dead insects exhibit rigor mortis (stiffening of the

muscles), which makes their appendages difficult to handle, and it is usually better to keep them in the killing bottle or in a hydrated atmosphere for 8 –24 h (depending on size and species) until they have relaxed (see below), rather than pin them immediately after death It should be noted that some large insects, espe-cially weevils, may take many hours to die in ethyl acetate vapors and a few insects do not freeze easily and thus may not be killed quickly in a normal household freezer

It is important to eviscerate (remove the gut and other internal organs of ) large insects or gravid females (especially cockroaches, grasshoppers, katydids, man-tids, stick-insects, and very large moths), otherwise the abdomens may rot and the surface of the specimens go greasy Evisceration, also called gutting, is best carried out by making a slit along the side of the abdomen (in the membrane between the terga and sterna) using fine, sharp scissors and removing the body contents with a pair of fine forceps A mixture of 3 parts talcum powder and 1 part boracic acid can be dusted into the body cavity, which in larger insects may be stuffed carefully with cotton wool

The best preparations are made by mounting insects

while they are fresh, and insects that have dried out

must be relaxed before they can be mounted Relaxing

involves placing the dry specimens in a water-saturated

atmosphere, preferably with a mold deterrent, for one

to several days depending on the size of the insects A

suitable relaxing box can be made by placing a wet

sponge or damp sand in the bottom of a plastic

con-tainer or a wide jar and closing the lid firmly Most

smaller insects will be relaxed within 24 h, but larger

specimens will take longer, during which time they

should be checked regularly to ensure they do not

become too wet

Pinning, staging, pointing, carding, spreading,

and setting

Specimens should be mounted only when they are

fully relaxed, i.e when their legs and wings are freely

movable, rather than stiff or dry and brittle All

dry-mounting methods use entomological macropins –

these are stainless steel pins, mostly 32– 40 mm long,

and come in a range of thicknesses and with either a

solid or a nylon head Never use dressmakers’ pins for

mounting insects; they are too short and too thick There

are three widely used methods for mounting insects

and the choice of the appropriate method depends on

the kind of insect and its size, as well as the purpose

of mounting For scientific and professional collections,

insects are either pinned directly with a macropin,

micropinned, or pointed, as follows

Direct pinning

This involves inserting a macropin, of appropriate

thickness for the insect’s size, directly through the

insect’s body; the correct position for the pin varies

among insect orders (Fig 17.2; section 17.2.4) and it

is important to place the pin in the suggested place

to avoid damaging structures that may be useful in

identification Specimens should be positioned about

three-quarters of the way up the pin with at least 7 mm

protruding above the insect to allow the mount to be

gripped below the pin head using entomological forceps

(which have a broad, truncate end) (Fig 17.3)

Speci-mens then are held in the desired positions on a piece

of polyethylene foam or a cork board until they dry,

which may take up to three weeks for large specimens

A desiccator or other artificial drying methods are

re-commended in humid climates, but oven temperature

should not rise above 35°C

Fig 17.2 Pin positions for representative insects: (a) larger beetles (Coleoptera); (b) grasshoppers, katydids, and crickets (Orthoptera); (c) larger flies (Diptera); (d) moths and butterflies (Lepidoptera); (e) wasps and sawflies (Hymenoptera); (f ) lacewings (Neuroptera); (g) dragonflies and damselflies (Odonata), lateral view; (h) bugs, cicadas, and leaf- and planthoppers (Hemiptera: Heteroptera, Cicadomorpha, and Fulgoromorpha)

Micropinning (staging or double mounting)

This is used for many small insects and involves pinning the insect with a micropin to a stage that is mounted on

a macropin (Fig 17.4a,b); micropins are very fine, headless, stainless steel pins, from 10 to 15 mm long,

Fig 17.3 Correct and incorrect pinning: (a) insect in lateral

view, correctly positioned; (b) too low on pin; (c) tilted on long

axis, instead of horizontal; (d) insect in front view, correctly

positioned; (e) too high on pin; (f ) body tilted laterally and

pin position incorrect Handling insect specimens with

entomological forceps: (g) placing specimen mount into foam

or cork; (h) removing mount from foam or cork ((g,h) After

Upton 1991.)

Fig 17.4 Micropinning with stage and cube mounts: (a)

a small bug (Hemiptera) on a stage mount, with position of pin in thorax as shown in Fig 17.2h; (b) moth (Lepidoptera)

on a stage mount, with position of pin in thorax as shown

in Fig 17.2d; (c) mosquito (Diptera: Culicidae) on a cube mount, with thorax impaled laterally; (d) black fly (Diptera: Simuliidae) on a cube mount, with thorax impaled laterally (After Upton 1991.)

and stagesare small square or rectangular strips of

white polyporus pith or synthetic equivalent The

micropins are inserted through the insect’s body in the

same positions as used in macropinning Small wasps

and moths are mounted with their bodies parallel to the

stage with the head facing away from the macropin,

whereas small beetles, bugs, and flies are pinned with

their bodies at right angles to the stage and to the left

of the macropin Some very small and delicate insects

that are difficult to pin, such as mosquitoes and other

small flies, are pinned to cube mounts; a cube of pith

is mounted on a macropin and a micropin is inserted

horizontally through the pith so that most of its length

protrudes, and the insect then is impaled ventrally or

laterally (Fig 17.4c,d)

Pointing

This is used for small insects that would be damaged by

pinning (Fig 17.5a) (but not for small moths because

the glue does not adhere well to scales, nor flies because

important structures are obscured), for very

sclerot-ized, small to medium-sized insects (especially weevils

and ants) (Fig 17.5b,c) whose cuticle is too hard to

pierce with a micropin, or for mounting small

speci-mens that are already dried Points are made from

small triangular pieces of white cardboard which either can be cut out with scissors or punched out using a special point punch Each point is mounted on a stout macropin that is inserted centrally near the base of the triangle and the insect is then glued to the tip of the point using a minute quantity of water-soluble glue, for example based on gum arabic The head of the insect should be to the right when the apex of the point is directed away from the person mounting For most very small insects, the tip of the point should contact the insect on the vertical side of the thorax below the wings Ants are glued to the upper apex of the point, and two or three points, each with an ant from the same nest, can be placed on one macropin For small insects with a sloping lateral thorax, such as beetles and bugs, the tip of the point can be bent downwards slightly before applying the glue to the upper apex of the point

Carding

For hobby collections or display purposes, insects (especially beetles) are sometimes carded, which involves gluing each specimen, usually by its venter, to

a rectangular piece of card through which a macropin passes (Fig 17.5d) Carding is not recommended for adult insects because structures on the underside are

Fig 17.5 Point mounts: (a) a small wasp; (b) a weevil; (c) an ant Carding: (d) a beetle glued to a card mount

(After Upton 1991.)

obscured by being glued to the card; however, carding

may be suitable for mounting exuviae, pupal cases,

puparia, or scale covers

Spreading and setting

It is important to display the wings, legs, and antennae

of many insects during mounting because features

used for identification are often on the appendages

Specimens with open wings and neatly arranged legs

and antennae also are more attractive in a collection

Spreading involves holding the appendages away

from the body while the specimens are drying Legs and

antennae can be held in semi-natural positions with

pins (Fig 17.6a) and the wings can be opened and held

out horizontally on a setting board using pieces of

tracing paper, cellophane, greaseproof paper, etc (Fig

17.6b) Setting boards can be constructed from

pieces of polyethylene foam or soft cork glued to sheets

of plywood or masonite; several boards with a range of

groove and board widths are needed to hold insects

of different body sizes and wingspans Insects must be

left to dry thoroughly before removing the pins and/or

setting paper, but it is essential to keep the collection data associated correctly with each specimen during drying A permanent data label must be placed on each macropin below the mounted insect (or its point or stage) after the specimen is removed from the drying

or setting board Sometimes two labels are used – an upper one for the collection data and a second, lower label for the taxonomic identification See section 17.2.5 for information on the data that should be recorded

17.2.2 Fixing and wet preservation

Most eggs, nymphs, larvae, pupae, puparia, and soft-bodied adults are preserved in liquid because drying usually causes them to shrivel and rot The most com-monly used preservative for the long-term storage of insects is ethanol(ethyl alcohol) mixed in various con-centrations (but usually 75 – 80%) with water How-ever, aphids and scale insects are often preserved in

lactic-alcohol, which is a mixture of 2 parts 95%

Fig 17.6 Spreading of appendages prior to drying of specimens: (a) a beetle pinned to a foam sheet showing the spread antennae and legs held with pins; (b) setting board with mantid and butterfly showing spread wings held in place by pinned setting paper ((b) After Upton 1991.)

ethanol and 1 part 75% lactic acid, because this liquid

prevents them from becoming brittle and facilitates

subsequent maceration of body tissue prior to slide

mounting Most immature insects will shrink, and

pig-mented ones will discolor if placed directly into ethanol

Immature and soft-bodied insects, as well as specimens

intended for study of internal structures, must first be

dropped alive into a fixativesolution prior to liquid

preservation All fixatives contain ethanol and glacial

acetic acid, in various concentrations, combined with

other liquids Fixatives containing formalin (40%

formaldehyde in water) should never be used for

speci-mens intended for slide mounting (as internal tissues

harden and will not macerate), but are ideal for

speci-mens intended for histological study Recipes for some

commonly employed fixatives are:

KAA– 2 parts glacial acetic acid, 10 parts 95% ethanol,

and 1 part kerosene (dye free)

Carnoy’s fluid– 1 part glacial acetic acid, 6 parts 95%

ethanol, and 3 parts chloroform

FAA– 1 part glacial acetic acid, 25 parts 95% ethanol,

20 parts water, and 5 parts formalin

Pampel’s fluid– 2– 4 parts glacial acetic acid, 15 parts

95% ethanol, 30 parts water, and 6 parts formalin

AGA– 1 part glacial acetic acid, 6 parts 95% ethanol,

4 parts water, and 1 part glycerol

Each specimen or collection should be stored in a

separ-ate glass vial or bottle that is sealed to prevent

evapora-tion The data label (section 17.2.5) should be inside

the vial to prevent its separation from the specimen

Vials can be stored in racks or, to provide greater

pro-tection against evaporation, they can be placed inside a

larger jar containing ethanol

17.2.3 Microscope slide mounting

The features that need to be seen for the identification of

many of the smaller insects (and their immature stages)

often can be viewed satisfactorily only under the higher

magnification of a compound microscope Specimens

must therefore be mounted either whole on glass

microscope slides or dissected before mounting

Fur-thermore, the discrimination of minute structures may

require the staining of the cuticle to differentiate the

various parts or the use of special microscope optics

such as phase- or interference-contrast microscopy

There is a wide choice of stainsand mounting media,

and the preparation methods largely depend on which

type of mountant is employed Mountantsare either

aqueous gum-chloral-based (e.g Hoyer’s mountant,

Berlese fluid) or resin-based (e.g Canada balsam,

Euparal) The former are more convenient for prepar-ing temporary mounts for some identification purposes but deteriorate (often irretrievably) over time, whereas the latter are more time-consuming to prepare but are permanent and thus are recommended for taxonomic specimens intended for long-term storage

Prior to slide mounting, the specimens generally are “cleared” by soaking in either alkaline solutions (e.g 10% potassium hydroxide (KOH) or 10% sodium hydroxide (NaOH)) or acidic solutions (e.g lactic acid

or lactophenol) to macerate and remove the body con-tents Hydroxide solutions are used where complete maceration of soft tissues is required and are most appropriate for specimens that are to be mounted in resin-based mountants In contrast, most gum-chloral mountants continue to clear specimens after mounting and thus gentler macerating agents can be used or, in some cases, very small insects can be mounted directly into the mountant without any prior clearing After hydroxide treatment, specimens must be washed in a weak acidic solution to halt the maceration Cleared specimens are mounted directly into gum-chloral mountants, but must be stained (if required) and dehydrated thoroughly prior to placing in resin-based mountants Dehydration involves successive washes in

a graded alcohol (usually ethanol) series with several changes in absolute alcohol A final wash in propan-2-ol(isopropyl alcohol) is recommended because this alcohol is hydrophilic and will remove all trace of water from the specimen If a specimen is to be stained (e.g in

acid fuchsinor chlorazol black E), then it is placed, prior to dehydration, in a small dish of stain for the length of time required to produce the desired depth of color

The last stage of mounting is to put a drop of the mountant centrally on a glass slide, place the specimen

in the liquid, and carefully lower a cover slip onto the preparation A small amount of mountant on the underside of the cover slip will help to reduce the likeli-hood of bubbles in the preparation The slides should

be maintained in the flat (horizontal) position during drying, which can be hastened in an oven at 40 – 45°C; slides prepared using aqueous mountants should be oven dried for only a few days but resin-based moun-tants may be left for several weeks (Canada balsam mounts may take many months to harden unless oven dried) If longer-term storage of gum-chloral slides is required, then they must be “ringed” with an