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British Culicoides
Contents | Introduction | Individual species pages | References
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Ceratopogonid midges are small nematocerous Diptera, the females of which have biting mouthparts and mandibles which work rather as a pair of scissors. They are mostly one to two millimetres long; the largest British species spans four millimetres or less. The body is stout and the wings are held flat and folded over the abdomen when at rest. They can easily be mistaken for members of the Chironomidae - indeed, they were formerly included in this family. They may be distinguished, however, by the short front legs (in the Chironomidae the front legs are usually much longer than the others), the wing venation and by the biting mouthparts (non-biting mouthparts in the Chironomidae). The wing venation
is simple with (usually) two radial cells. In most species the wings are clear, without markings; but many species of Culicoides (and of some species of other genera) have easily recognisable patterned wings (some Chironomidae have patterned wings, but then the venation is different). A good formal diagnosis of the family is given in Downes & Wirth (1981), and a key to the British genera of Ceratopogonidae is provided in Boorman & Rowland (1988)
It is important to realise that the British fauna is only a part of the wider Palaearctic fauna, and that the British fauna cannot be studied in isolation. Much useful information is to be found in literature concerned with other parts of the Palaearctic, and species may also be referred to by other names. These are the reasons for the inclusion of many “foreign” references and synonyms. Forty seven valid species have been noted in Britain, but others undoubtedly remain to be discovered, either having been overlooked, or may yet appear here as a result of global warming and consequent immigration from the Continent.
Some 5500 species of Ceratopogonidae are recognised worldwide, belonging to five subfamilies and about 125 genera, of which almost 25 are represented by fossil species (Borkent & Wirth 1997). Some authors have argues that the genus Leptoconops and the related fossil genera represent a distinct family, the Leptoconopidae (Downes 1977 ), but these genera are usually included in a separate subfamily, the Leptoconopinae. Leptoconops midges do not occur in the U.K, although several species are found in the warmer parts of the Palaearctic region - for instance, North Africa. Culicoides belong to the subfamily Ceratopogoninae. This includes six tribes including the tribe Culicoidini. Only one genus of this tribe occurs in the U.K.; Culicoides.
The genus Culicoides was erected by Latreille in 1809, although the first description of the life history of one of these biting flies was that of the Rev. W. Derham, the Rector of Upminster, in 1713; he bestowed on them the name Culex minimus nigricans maculatus sanguisuga. From this name and his description of them biting horses in numbers, it may be supposed that he was writing of either Culicoides pulicaris or Culicoides nubeculosus.
Certain taxonomic conventions should be observed. Generic names always begin with an upper case letter and specific names are always lower case. Authors names are sometimes abbreviated (for example, Mg. For Meigen) but this may lead to confusion and names are best quoted in full. The original assignment of a species to a particular genus may not always be correct. When, after further work, a species has been transferred to a genus other than that in which it was originally described, the author’s name is given in parentheses (for example, Culicoides obsoletus (Meigen) was originally described in the genus Ceratopogon).
A species may described from a single specimen or a series of specimens. In this case it is usual to select one specimen as the holotype; others studied at the same time and included as part of the type series are known as paratypes. Paratypes are always of lesser status than the holotype, since it is possible that they may have been incorrectly assigned. The holotype may be of either sex, the choice being that of the author; it is usually a specimen which displays the diagnostic features to the best advantage. The term allotype was used formerly for a specimen in the type series of the opposite sex to the holotype; the term is no longer used and should be avoided. Where a species is described from a series of specimens but no single one is selected as the holotype, each member of the series is termed a syntype (formerly cotype). In museum collections holotypes are usually denoted by a red label; paratypes have yellow labels.
It was formerly the practice to preserve Ceratopogonidae dry, either pinned or on card points. This has led over the years to many species being represented by bare pins. When the holotype has been lost (as in the case where much material was lost in a disastrous fire in the Hungarian National Museum) a neotype may be described, preferably from the same locality and of the same sex as the original holotype. This should normally be done only where it is certain that the type has been lost; otherwise it is always possible that the specimen may be found in another collection (this frequently has been the case in many of Kieffer’s species).
There are many cases where a species has been described under two or more different names (for instance, see C. circumscriptus or C. puncticollis). In such cases, the law of priority applies and the name under which the species was first described is held to be the correct one; all other names are then referred to as junior synonyms. In rare cases the Committee for Zoological Nomenclature may rule that an alternative should be used, for instance in the case of Culicoides puncticollis, which was first described under the name of C. algecirensis. Here, the use of the latter name could lead to confusion in a species of potential veterinary importance and the commonly used name has been re-instated.
Several attempts have been made to place the species worldwide into subgenera; these have been only partially successful and the subgeneric classification of Culicoides remains a subject of debate. Remm (1988) recognised seven Palaearctic subgenera; Glukhova (1989) recognised eleven, members of four of which have not been recorded from Britain. I have followed Glukhova's analysis; she has rightly pointed out the importance of study of the early stages as well as of adults. I have not included descriptions of known larvae here, since the list would not only be incomplete but it would add greatly to the length of this review. Also, since the whole subject is still undecided, I have chosen not to give a key to subgenera. One of the main stumbling-blocks to the wide acceptance of subgenera has been the status of the subgenus Oecacta, which is usually used as a "catch-all" to include those species which do not fit in elsewhere. The genus Oecacta was erected by Poey (1853) for Culicoides furens. This species is closely related to the schultzei-kingi-oxystoma group, for which Glukhova (1977) erected the subgenus Remmia. If one accepts, as is most probable, that furens and schultzei are related, then Remmia becomes a junior synonym of Oecacta. The majority of species included in Oecacta are clearly not related to the schultzei group and must therefore be placed elsewhere (Boorman 1988, Cornet & Brunhes 1994). For the moment, however, they are left as most commonly accepted.
The following are the main characters separating the subgenera known to be represented in Britain, and will go some way towards giving a clue to the identity of individual species. The species are grouped accordingly (and alphabetically) on the Contents page, and the links below lead to individual pages for each species, also showing wing photographs and diagrams of male genitalia for identification.
Avaritia (4 species)
Second radial cell pale, wing markings often vague and ill-defined; third segment of palp with a single small circular sensory pit; female antenna with sensilla on 3, 11-15; two spermathecae; ninth tergite in the male without apicolateral processes.
Beltranmyia (3 species)
Second radial cell dark, wings dark with pale circular markings; female antenna with sensilla on segments 3-14; one spermatheca.
Culicoides s.s. (7 species)
Second radial cell pale; wing pale with darker markings; third segment of palp with scattered sensilla but without a clearly defined single sensory pit; female antennae with sensilla on segments 3, 11-15; two spermathecae.
Monoculicoides (5 species)
Second radial cell dark; wings pale with darker markings or only the second radial cell darkened; female antennae with sensilla on segments 3, 8-10; one spermatheca.
Oecacta (19 species)
Second radial cell dark or pale; wing markings various; female antennal sensilla various; two spermathecae.
Silvaticulicoides (5 species)
Second radial cell dark; wing with at least two pale spots, one over R-M and the other just beyond the second radial cell; female antennae with sensilla on segments 3, 11-15; two spermathecae.
Wirthomyia (3 species)
Wings without markings; female antennae with sensilla on segments 3-10 or 3-14; two spermathecae.
Unplaced (C. cameroni only)
Wings unmarked; female antennae with sensilla only on segment 3; male genitalia distinctive.
The eggs are laid singly or in small batches on any debris in the breeding site; all Culicoides larvae need a certain amount of free water to complete their life cycle. Some species breed in wet leaf litter; others in habitats ranging from mud through tree holes to the banks of streams or among floating vegetation on lakes. There are four larval instars. The majority of species overwinter as larvae; a very few as eggs but apparently none as pupae. Pupae are formed amongst litter or float on the surface of free water in the breeding site. The egg stage lasts usually for three days to a week; but Jobling (1953) found that the eggs stage in C. vexans the egg stage may last over four months, while the larval stage may last for six months. Usually, however, the larval stage lasts for two weeks to a month or so.
The majority of species are univoltine, but some have several generations through the year, depending on local conditions.
Male midges may be found feeding on nectar in flowers; a large proportion of adults of both sexes (with the exception of freshly emerged adults) test positive for reducing sugars in the anthrone test (van Handel 1972). Male midges do not take blood, but most females require a blood meal to mature a batch of eggs. Some species, for instance C. riethi, C. impunctatus and C. circumscriptus mature a first batch of eggs autogenously but then take a blood meal for subsequent egg batches. The gonotrophic cycle usually occupies from one to three weeks, depending upon the ambient temperature. Adult midges may live for a week to a month or more; a female C. obsoletus survived for three months in a cool insectary (personal observation). Most midges disappear after the first frosts of winter, but catches throughout the winter months have yielded the occasional C. obsoletus from December to February in the south of England, suggesting that they may occasionally survive the winter..
Midges have been age graded by dissection and observation of follicular relics, but this procedure is very time consuming and technically difficult. Changes in the ovaries, and ovarian cycles in both Culicoides and Leptoconops have been described by Linley (1965, 1965 and 1966). Dyce (1969) observed that parous females of many species develop a cherry-red pigment in the subcutaneous cells of the abdomen, and this is easily seen in many species in fluid under a stereomicroscope. This observation gives a method for the easy separation of nulliparous and parous midges. Birley & Boorman (1982 ) have shown that estimates of longevity and gonotrophic cycle length can be made by observing the relative proportions of nulliparous and parous females using this method over a period of over eleven successive days, providing that certain conditions are met.
Culicoides have been incriminated as vectors of a wide variety of pathogens; viruses, filariae and a range of protozoan blood parasites. In the British Isles they have been shown to be a vector of the filaria Onchocerca cervicalis of horses, while at least one species is a potential vector of blue tongue virus of sheep and cattle in southern and southeastern Europe. Their role as virus vectors has been reviewed recently by Mellor et al. (2000).
Abroad, and particularly in the tropics, Culicoides may be an important biting pest; in Britain, Culicoides impunctatus is an important biting pest in Scotland where its effect on tourism and forestry is considerable (Hendry 1996)
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, 1979).
Several species of Culicoides have been maintained as laboratory colonies; for example in Britain, C. nubeculosus and C. riethi. (Boorman, 1974).
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 blue tongue 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.
CAUTION Alcohol-phenol is VERY CAUSTIC and must be handled with very great care. When handling it or when slide-mounting NEVER allow ones fingers to touch ones face, particularly the eyes. Wash the hands well after handling phenol-balsam or phenol-alcohol.
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.
The microscope, whether a steromicroscope or a compound microscope, is a precision scientific instrument and should always be treated as such. When not in use, even for short periods, always cover the instrument with some form of dust cover. Lenses, in particular eyepieces, should be kept clean and free of dust. Remove dust by blowing gently or brushing gently with a fine soft brush (as sold for cleaning camera lenses). Oily smudges should be removed by wiping gently with a soft cloth or piece of chamois leather, as used for cleaning spectacles. If necessary, moisten the cloth with a little xylene or fluid sold specifically for cleaning optical lenses; never use spirit as this may dissolve the cement used to fix the lenses in place.
It is great importance that the microscope should be adjusted correctly to suit the user. This applies to both types of instrument. The eyepieces should be correctly adjusted for focus for both eyes, and the separation of the eyepieces set for the user. These simple adjustments can make a great deal of difference, not only to discerning fine details, but also to avoid eye strain.
Compound microscopes should always be set up correctly and this should be checked at the beginning of each session. Firstly, check the focus of the eyepieces by placing a specimen on the stage and examining under, say, a x10 objective. Adjust the eye separation.
Most modern instruments will have two adjustable diaphragms, one between the light source and the condenser (the field diaphragm) and the other on the substage condenser itself (the iris diaphragm). Close both and view the specimen under a x10 objective. Focus the field diaphragm by adjusting the substage condenser and centre it by adjusting the condenser adjusting screws. Open the field diaphragm so that the field is completely lit. Open the iris diaphragm so that it fills about two thirds of the field (remove one eyepiece and look down the barrel of the eyepiece to see this). The instrument is now set up for the best results.
Do not control the amount of light on the specimen by adjusting the iris diaphragm; use the lamp transformer for this. Stopping down the iris to two thirds as above will resolve the greatest detail in the specimen; stopping down further decreases the resolution although it may apparently make some structures (often the antennal sensilla) easier to see.
Dark field illumination is of great value in making out the presence of light and dark spots on the wings, but bear in mind that pale spots will appear as dark, and vice versa. Some (but not all) modern condensers will have a dark field setting; use a low power objective and open the iris diaphragm fully. In some cases a false dark field effect can be obtained by choosing a low power phase objective and a mis-matched phase ring, but this is not as effective as a proper dark field condenser. The slide should be clean and dust free, but beware if the balsam is not quite dry!
Phase contrast can be very useful to see details of wing venation and sometimes also the presence of antennal sensilla. Always check to make sure that the microscope is set up for phase illumination in accordance with the manufacturers instructions, and particularly if the phase rings are accurately centred.
With practice, some adult Culicoides may be identified under a hand lens or a stereomicroscope, but the majority must be slide-mounted. The characters of most value are:
A summary of the characters used is given below.
The eyes are sometimes useful; in some species they are separated, in other joined (contiguous). Occasionally the eyes bear minute bristles or setae between the facets - that is, the eyes are described as “hairy”. This is most easily seen in specimens which have been treated with caustic potash and cleared, but may also be seen by looking carefully at the edge of the eye.
The form of the palps, and particularly of the third segment, and the length of the proboscis, provide two useful diagnostic features. The third segment is often swollen, and the “palp ratio” is the length of segment 3 divided by its greatest breadth. The P/H ratio is the distance from the end of the labrum-epipharynx to the tormae divided by the distance of the tormae to the base of the interocular seta. The third palpal segment bears a cluster of sensilla or a sensory pit, the form of which is useful in separating some species.
The antennae are of considerable diagnostic importance. They consist of fifteen segments: a scape, a pedicel, and thirteen flagellar segments. This is the origin of the antennal segments being referred to as 3 - 15, or III to XV; it is more accurate to refer to “flagellar segments” or “flagellomeres”. The antennal ratio a/b is found by dividing the combined length five apical segments (i.e. 11-15) by the combined length of the basal eight (i.e. segments 3-10). The various segments bear a variety of sensilla, the most important of which are the sensilla coeloconica. These are typically minute pits surrounded by short setae and are usually found towards the apex of the segment; they vary in number from one to several and may be present or absent on individual segments. All Culicoides have sensilla coeloconica on segment 3; and the distribution of the sensilla in the female (for instance, they may be present on 3-15, or 3, 11-15 only, or 3,5,7,9,11-15 only, or other combinations.) is often diagnostic, may vary in some species.
The antennal segments bear other sensilla, the sensilla trichodea, sensilla basiconica, sensilla ampullacea and sensilla chaetica (Cornet 1974). They have been found useful in distinguishing some species, but are not generally of importance. The sensilla trichodea (soies transparentes of some French authors) are useful in separating some members of the obsoletus group, but not in the U.K. The antennae of male midges also bear sensilla coeloconica, but these have a different distribution to the female. They have not been widely studied.
The mesonotum may be plain, unmarked, or in some species may bear prominent darker markings. These are most easily seen in dry specimens although the mesonotum may be distorted; but if the mesonotum is separated during slide-mounting and mounted flat with the dorsal side uppermost any markings may be made out under the microscope. The legs occasionally bear darker bands or patches but are not usually of great value in identification of Culicoides.
The wings are of primary importance. The venation (fig. 3) is simple with usually two radial cells. A figure of a typical Culicoides wing is given in figure 1, together with the names of the veins (abbreviations beginning with upper case letters) and cells (abbreviations beginning with lower case letters). Older descriptions may employ different names for cells and veins; Kremer (1966) gives a table with older names. Many Culicoides have the wings patterned with pale or dark spots, and the disposition and extent of these separates the various species. The wings also give a useful indication of relative size. The length of the wing (l) is measured from the basal arculus to the wing tip, and the length of the costa (c) from the arculus to the end of the second radial cell. The costal ratio is found by the ratio c/l, although this ratio is of limited use in the genus Culicoides.
The number of functional spermathecae in the female is an important diagnostic aid. Members of the subgenera Beltranmyia and Monoculicoides have a single spermatheca. Most species have two functional spermathecae and a rudimentary third. No British species have three spermathecae (the occasional aberrant individual may have three) although members of the genera Trithecoides and Pontoculicoides (found abroad including other parts of Europe) have three.
The genitalia of the male provide one of the most important diagnostic features. The shape of the ninth tergite and particularly its posterior border, the absence or presence of lateral processes, the shape of the aedeagus and of the parameres are important. The ninth sternite may be emarginate and the membrane covering the sternite may have numerous minute spicules, or may be bare. The shape of the ninth sternite is of particular value in differentiating the members of the obsoletus group. The ventral roots of the basistyles also provide clues; in some species they are long and tapering, in others they are foot-shaped.
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