Here we give only a brief account of methods of collecting, preserving and making slide preparations of aphids. For more detailed techniques of aphid study consult van Emden (1972) or Minks & Harrewijn (1988).



Aphids on trees and shrubs may be collected by sweeping or beating, or simply by careful searching of appropriate parts of the plant. Beating onto a tray or card held underneath the branch may be useful for free-living aphids on leaves and shoots of trees or shrubs, especially Lachninae on conifers, but it is not very efficient for active, winged aphids such as many Calaphidinae, adults of which will fly as soon as disturbed leaving only the immatures and damaged adults to fall onto the beating tray. Beating cannot be used to collect aphids that live in wax wool such as Adelgidae, and is obviously useless for those making galls or feeding on the trunk or large branches.

It is best, if at all possible, to examine the plant carefully and try to find the aphids in situ, so that a representative sample of all available morphs and developmental stages can be collected and information gained about the feeding site and the size and appearance in life of the colony, if one is formed. Aphids inhabiting the undersides of leaves can often be observed in silhouette by looking up through a leaf against the sun. Ant-attended aphids, such as Thelaxinae, Chaitophorinae and many Lachninae, can often be detected by looking for ants moving up the trunk of the tree. Large numbers of active flies and wasps may also indicate presence of honeydew-producing insects.

Collecting aphids on herbaceous plants is simply a matter of careful examination of all parts of the plants, not neglecting to look on the undersides of leaves, and in the inflorescences. Aphids colonising the basal parts of plants or roots are often detectable by the presence of attendant ants, or may be covered by earthen shelters built by the ants. Some aphids tend to live on etiolated stems or runners growing in darkness under stones. Some are highly cryptic and may only be revealed by use of a beating tray. Beating may also be the best method of collecting certain aphids that escape by falling off the plant  at the least disturbance. Sweeping herbaceous vegetation is not recommended as it is rarely possible to ensure that only one species of plant is being swept at a time, and it is always important to know the identity of the host whenever possible.  

It is usually considered best to bring the aphids back to the laboratory alive on a piece of the host plant, rather than to collect them directly into preservative. Large polythene or glass tubes stoppered with cotton wool, polystyrene sandwich boxes or polythene bags are suitable containers. A piece of tissue paper should be included in each container to soak up excess moisture, and plenty of air should be included in a polythene bag before it is tied up. In warm weather it is preferable to place samples in an insulated cool bag or box with ice blocks, or a large precooled vacuum flask, especially if they are going to be left in a car for even a short time. The reason for keeping specimens alive is that colonies often consist of mainly immature individuals, and the proportion of adults in the sample can be increased by keeping the aphids for a few days in a cool place before preserving them. Adults, especially alatae, should be left for one to two days after they have reached maturity in order to develop their full pigmentation. This also provides a method of rearing out any parasitoids. It is important, of course, to make sure first that no predators are included in the container!

Another method of collecting tree-living aphids is to look for their eggs on the twigs, branches or trunk in late summer or early spring. Pick a sunny day as many aphid eggs are shiny black and much more conspicuous in sunlight. Any twigs found to have eggs on them can be cut off, brought back to the laboratory and placed in water; the eggs are then likely to hatch quite quickly, and the hatching fundatrices can often be reared to adult on the swelling and breaking buds and young shoots, although to rear a second generation fresh twigs will almost certainly be required.

It is preferable to note down the fullest possible collection data at the time of collection, including host plant, locality and date, and it is also important to note biological information such as the colour of the aphids in life (both adults and immatures), feeding site, whether the aphids occur singly or form a colony, and whether or not there is ant-attendance. It is probably best to keep a field notebook for this purpose, and to give each sample a collection number which is written on a slip of paper and inserted in the container with the specimens. If the specific identity of the plant is unknown or uncertain then leaves and, if possible, flowers or fruits should be collected for examination by a botanist; notes should also be made on the growth habit of the plant, and the form and texture of the bark in the case of trees. A fine sable hair brush for handling aphids, a good-quality hand lens and a small pair of secateurs are essential field equipment.


Aphids for morphological examination should be preserved in tightly stoppered tubes filled with 80-90 ethanol. For prolonged storage one volume of 75 w/w lactic acid may be added after a few days to every two volumes of alcohol containing specimens, and the tubes plugged with cotton wool and kept under alcohol or on a cushion of cotton wool soaked in alcohol, in an air-tight glass jar.

Maceration to remove the soft tissues of the specimen prior to mounting is best carried out with the specimen tubes in a water bath kept near boiling point, or on a dry-block heater. The following stages are involved:

1. Gently boil the specimens in 95% ethanol for 5 min.

2. Transfer the specimens to 50% ethanol in a watch-glass, sort specimens for mounting and prick the abdomens with a sharp needle of fine entomological pin.

3. Decant or pipette off ethanol, add about 1 cm depth of 10% potassium hydroxide (KOH) solution, simmer for 3-5 min. Check that body contents have become transparent; if not, leave for an extra 2-3 min, or if necessary expel some of the body contents by gently squeezing the abdomen with forceps.

4. Decant or pipette off KOH solution and wash the specimens free of all KOH using 5-6 changes of distilled or deionized water, leaving them to soak for at least 5 min each time.

The water-based Berlese mountant has frequently been used in the past by aphid workers, but balsam mounts are recommended because of their proven permanence and resistance to a wide range of climatic conditions. The macerated specimens need to be totally dehydrated and cleared before mounting in Canada balsam. This can be done most simply using Martin's (1983) method:

5. Remove distilled water, add 1 cm depth of glacial acetic acid and leave for 2-3 min. Pipette off and repeat with fresh glacial acetic acid. Pipette off.

6. Add clove oil as clearing agent (specimens will float). Leave for 10-20 min until specimens are clear.

7. Transfer 1-2 aphids to a drop of fairly thin Canada balsam on a clean microscope slide and quickly arrange them with body untwisted, dorsal side uppermost and appendages spread out.

8. Dip a clean 16 mm round coverslip in xylene and immediately lower it carefully onto the specimens displayed in the drop so as to spread the mountant evenly without trapping air bubbles.

9. Dry the slide horizontally in an oven at 50°C for about 1 week.


For a more detailed account of mounting procedures for aphids and other small insects, and for advice on remounting of deteriorating aphid slides, see Brown & De Boise (2004).

For those who do not have ready access to Canada balsam or xylene, the following method using PVA glue as the mountant can provide slide-mounted specimens of excellent clarity, enabling specimens to be taken through the keys on this website, although the preparations do not have the proven permanence of balsam mounts:-

1-4.  As 1-4 above.

5. Transfer specimens from final wash to 20% PVA clear glue (e.g. “Pritt in a Tube”, available from stationers and art shops) in distilled water (shake the liquid thoroughly and leave for an hour before use).

6. Transfer specimens to a watch-glass, and place one in a drop of PVA clear glue squeezed straight from the tube onto the centre of a clean microscope slide. Alternatively, for a better refractive index and longer-lasting result use PVA/borax/glycerol mountant (“PVA-G”; see Dioni 2003). To make this add 6 ml of glycerol to 10 ml of PVA clear glue, then add 4 ml of distilled water saturated with borax, mix well and leave for at least 24 hours, with occasional gentle warming until the liquid is clear of air bubbles. (Note: a great mass of small air bubbles is initially produced; these same ingredients are used to produce artificial “slime”!). Decant the clear mountant into a fresh container before use.

7.  Arrange the specimen quickly as in point 7 above. Apply a clean 16 mm round cover-slip, lower it carefully and press gently with forceps to spread mountant evenly without trapping air bubbles. Leave to dry overnight. If PVA is used as a mountant on its own then the cover-slip will need to be ringed after 24 hours with nail varnish to prevent ingress of air. The PVA/borax/glycerol mountant does not require sealing, but ringing with nail varnish will help to keep the cover-slip in place.

Additional notes

   If the rostrum is short and the thorax is very dark, as is often the case in alate specimens, try to displace the rostrum to one side so that the last rostral segment (R IV+V) is not obscured by the thorax. On the other hand, if the rostrum is long or the thorax pale then the rostrum is best placed ventrally along the midline, so that its length and the shape of R IV+V can be readily determined.

   Characters of the embryos contained within the maternal body, especially the number, size and arrangement of dorsal hairs, are often used in species identification, e.g. of alatae of Eriosomatinae and Hormaphidinae emerging from galls and of certain Calaphidinae. These characters can be observed through the maternal cuticle in well-prepared specimens, but are better displayed if a small incision is made in the abdomen and a few embryos are carefully squeezed out into the mounting medium before putting on the coverslip.


Aphid workers have found that thick card labels, glued to the slide with an impact adhesive, help to protect the coverslip and enable slides to be stacked vertically while awaiting attention. It is advisable to standardize the labelling; the system most frequently used is shown in fig. 135. Slides may be stored horizontally in trays, or vertically in slotted drawers or boxes. For larger collections a compact and versatile system involves vertical storage of slides in individual envelopes made of paper or cellulose acetate, with interspersed tab cards providing the collection with an integral index (Eastop 1985).