Studying Culicoides

Culicoides adults may be reared from mud or debris collected from potential breeding sites. Samples are kept in glass jars must be watched carefully as newly emerged adults are easily trapped in drops of condensation.

Third or fourth instar larvae may be separated from mud or other debris by flotation using magnesium sulphate or, with less chance of damaging the larvae, strong sucrose solution. Larvae float to the surface and may be picked off with a pipette, transferred to other mud or debris in small tubes and can often be successfully hatched.

In some cases it may be possible to induce gravid females to lay eggs if confined in glass tubes with damp filter paper or cotton wool; alternatively gravid females may be induced to lay mature eggs by lightly anaesthetising them and severing the head (Linley 1965). In many cases such eggs will prove to be infertile, but if the eggs turn from white to black some fifteen minutes or so after laying there is a good chance that they will hatch.

Newly hatched larvae can be placed on mud taken from a breeding site but are almost impossible to see and are rarely reared successfully. Small larvae may be fed on small nematodes on an agar substrate; this has the advantage that they may easily be seen and is occasionally successful (Linley 1979).

Several species of Culicoides have been maintained as laboratory colonies; for example in Britain, C. nubeculosus and C. riethi (Boorman, 1974). [click here for information on the Pirbright colonies]

There is a large amount of literature concerning the use of bait animals for collecting biting insects, particularly mosquitoes. Most of the methods used can be used for catching Culicoides providing any nets used have a sufficiently small mesh, and can yield useful information on their habits. They can also be a useful source of blood-fed insects if an attempt is to be made to colonise a species. The site of biting can also be investigated in this way; and the technique has ben used to reveal different biting sites in C. pulicaris and C. punctatus (Campbell & Pelham-Clinton 1960).

A fine mesh net may be used to collect Culicoides coming to bite or swarming; for example, this is a good way to collect C. heliophilus flying over heather during the day. Some species are attracted to the flowers of Umbelliferae and may be swept for during the day or evening, although sweeping vegetation is usually more successful in catching other genera of Ceratopogonidae.

Most, if not all, species of Culicoides are attracted to light traps and this is one of the most productive methods for general surveys of midge fauna. A large number of designs have been published; the choice depends largely on the facilities available; for instance, whether mains power is available or batteries must be used. In general, the brighter the light source, the more insects will be attracted. It should be remembered that the brighter the light source the more insects will be attracted, and the more time consuming it will be to sort the catch. Black light or mercury vapour lamps are particularly effective; but while small black light tubes can be battery powered, mercury vapour lamps will need mains power. An inverter to convert 12-volts to 120 or 240 volts can be used to drive a mercury vapour lamp from a car battery. If mains power is used, particular care must be taken to avoid accident, and the use of a residual current device or earth leakage trip is essential. At its simplest, a light trap can be made by suspending a torch bulb over a bowl containing water to which a little detergent has been added.

Insects may be collected dry, in a fine mesh cage, or wet into alcohol, or into water or saline (0.9%) with a little detergent to wet the insects. Traps should be emptied first thing the next day, as the insects quickly deteriorate in warm weather if left for long periods, resulting in a foul-smelling thick soup of wings and legs. If the catch cannot be recovered immediately, a little formalin (2%) added to the water will prevent too rapid a decomposition, but higher concentrations may have a repellent effect.

The choice of method for storing specimens depends on the purpose for which they are collected. If they are to be used for virus isolation or for DNA studies then the use of preservatives such as formalin (and possibly alcohol) must be avoided; depending on the method to be employed, freezing in liquid nitrogen may be required. Samples for the isolation of bluetongue or African horse sickness viruses cannot be stored at -20 degrees as this will result in the loss of virus. Storage in dry ice may also result in virus loss due to absorption of CO2 and lowering of the pH. The remarks which follow assume that the object is a taxonomic or faunistic study.

Again, the choice may be dictated by the method of collection. Insects collected dry may be stored dry in the cold in vials or mounted dry for examination. Midges were commonly pinned with fine stainless steel pins or mounted on card points, and stored dry in cabinet drawers. Many such specimens ended their useful life as bare pins (dried pinned midges are very brittle) or as a leg or two on a card point; and unless there are good reasons dry storage should be avoided.

Insects collected in saline (normal saline, 0.95%) with a little detergent added to wet the specimens, water or weak detergent may be preserved either in alcohol or formalin. Alcohol (70% ethyl alcohol or isopropyl alcohol) is a good general fixative, and more pleasant to use than formalin. It has the disadvantage that specimens tend to darken to a uniform brown over years and become brittle, making subsequent manipulation difficult. Formalin (1% or 2% commercial formalin solution, which is a 40% solution of formaldehyde in water) is unpleasant to use in greater strength (5% to 10%) but lower dilutions are perfectly adequate as a preservative provided that the catch has been well fixed before final storage. 2% dilution is perfectly adequate if a small number of insects are being stored in a tube or vial. Formalin has many advantages and is the preservative of choice for Ceratopogonidae. The colours of insects are retained (but formalin-preserved insects should be kept in the dark since light will rapidly bleach them) and the insects remain flexible and easy to dissect. Formalin can also be carried as a strong solution and diluted with water as required.

Whether alcohol or formalin is used for storage, bottles and vials may dry up over a period, leaving the insects useless for study. The addition of 1% glycerol to either alcohol or formalin will prevent such loss; as the preservative evaporates the relative glycerol concentration rises and the tubes will not dry out completely. Insects are left in a thin film of glycerol and can easily be re-wetted.

The preparation of slide mounts of adults is often essential for accurate identification, and is also desirable for the preservation of voucher specimens.

It is usually unnecessary to treat Culicoides with caustic potash before mounting; after clearing sufficient detail is easily visible. Very brittle specimens may need a brief soaking in dilute (5%) caustic, but the time should be kept to a minimum and heating should be avoided.

Various media have been used for the preparation of slides. In general, gum-chloral media (Berlese medium, Swann’s medium) are useful for temporary mounts where a quick answer is needed. Such media should never be used for permanent storage. They are always liable to drying out, resulting in shrinkage and possible damage to the specimen. Also, some formulations containing glucose are apt to turn black over the years, with destruction of the specimen within. Their main advantages are speed in preparation and that specimens can readily be removed from slides by soaking in water.

The medium of choice is undoubtedly Canada balsam. This is the only medium which has proved to be “permanent” over many years. Its chief disadvantages are that the methods are time-consuming, that slides take a long time to dry (many weeks or months) and that the balsam will darken somewhat over the years. However, specimens can always be soaked off with xylene or alcoholic phenol and remounted, although such specimens are often rather brittle. Wirth & Marston (1968) described a method of preparing permanent mounts using phenol-balsam; this has been adopted by many workers and is widely used.

The Canada balsam should be that described as “natural and filtered”; it is very thick and often will not pour unless the bottle is warmed cautiously with hot water. Balsam dissolved in xylene as commonly sold for microscopy is not suitable. Thick balsam should be thinned in alcohol-phenol, which is a saturated solution of phenol crystals in absolute ethyl alcohol. To prepare this add a very small amount of alcohol (10ml at a time) to a bottle of phenol crystals. The liquid will get very cold and should be allowed to warm up before adding more alcohol. Always aim to leave a thick layer of undissolved crystals at the bottom of the bottle.

Phenol-alcohol should be kept in a dark bottle away from light, as should diluted balsam as it will rapidly darken. For best results, dilute as small quantity of balsam at a time and do not store it for long periods. The amount of phenol-alcohol needed to dilute the balsam is a matter of personal preference; thicker solutions will not spread so easily on slides and specimens may prove easier to dissect. Only experience and personal preference can decide this.

Midges collected in either 70% alcohol or in formalin may be transferred directly to alcohol - phenol in a glass vial and will clear in around four hours, but they may be left in phenol for up to three weeks. Place four drops of phenol - balsam separately on the centre of a microscope slide and transfer the midge carefully to the lower left drop. Under a stereomicroscope at 12-15x magnification, remove the head remove the head and place it in the upper right drop. With cutting needles separate the wings from the thorax and place one or preferably both wings in the bottom right drop. With two cutting needles orient the thorax so that it lies on its side and slice off the mesonotum together with the scutellum, taking care not to cut the legs. Leave the mesonotum with the rest of the thorax and legs in the bottom left drop. Remove the abdomen and place it in the upper left hand drop of balsam.

Using 7mm square (approximately) cover slips cover the head, abdomen, wings and thorax. Before covering, orient the head so that it lies front side up with the antennae spread out: orient the abdomen ventral side uppermost; make sure the wings lie flat and the mesonotum is dorsal side uppermost.

It will be found helpful if the microscope stage is marked so that the slide is placed centrally, and mark the positions that the four drops of balsam should occupy. This will ensure that the parts can easily be found in the same place when successive slides are examined.

7mm. square cover slips are not generally available but may be cut from standard 22mm square cover slips using a glass-writing diamond (from a supplier of laboratory equipment) and a straight edge (a microscope slide). Place the cover slip on a hard dark surface with the light coming from in front, preferably from a window. Mark the cover slip, using gentle pressure and holding the diamond cutter vertically, with three scratches in each direction, making a total of nine small cover slips. Some trial and error is needed to obtain the right angle of the diamond and the right amount of pressure to mark the glass. Finally, separate the nine small slips using gentle pressure.

Suitable cutting needles can be made in two ways. “Hagedorn” surgical needles are flat broad needles with a single oblique cutting edge. They are obtainable from surgical instrument suppliers and come in a range of sizes. Small Hagedorn needles may be mounted using “Araldite” in 18-gauge hypodermic needles cut to 1 inch long with a triangular needle file. More easily, a range of different size needles may be made by flattening the end of standard hypodermic needles using a hammer while holding the needle on a flat piece of steel, holding the needle with the bevelled edge uppermost. This results in an arrow-shaped needle with two cutting edges. When it becomes blunted with use, discard it and make another. The needles may be mounted on a 1ml disposable hypodermic syringe; the plunger of the syringe (without the plastic seal) makes a convenient balsam dropper. Also needed will be a pair or two of very fine pointed forceps and also, if possible, a pair of fine point forceps with curved tips. These latter are very useful for picking up numbers of midges. Lastly, a hand lens with x8 or x10 magnification will be found useful.

As a working rule, never lend forceps or mounting needles to colleagues. Let them make or get their own. Needles or forceps with bent tips are useless.

If it is worth taking the trouble to make a slide of a midge, it is worth keeping it for future reference. It is very important to label slides adequately; a scribbled note with a felt tip pen is not acceptable. Slides should have one or preferably two 25mm labels, one at each end, to carry locality data, date of capture, collection method and captor, together with any identification and other notes (for example, sensilla distribution). Data should be written with a fine drawing pen using waterproof ink. Labels are best attached with a water soluble adhesive; ordinary PVA woodworking adhesive is ideal. Slides may be stored flat or stacked on their sides; in the latter case care must be taken that the slides do not rub onto each other. This may be accomplished by attaching a narrow strip of card at each end, or by using thin card for the slide labels themselves.

If the slide collection is of any size, the slides should be numbered to allow reference to a particular specimen. The details may then be entered in a computer database.

In museum collections, it is a convention to attach a red label to a holotype, yellow for a paratype, blue for a syntype and purple for a lectotype.