Adult biting midges may collected by any of the following techniques:
1. Sweeping vegetation, especially in wet habitats, and aspirating the adults from the net. This provides a good method to sample diverse habitats.
2. Light trap. Although many specialized light traps exist that work very well, a good sample may even be obtained at a lit window pane when there is good surrounding habitat. Ultraviolet light works well to attract adult midges but regular inflorescent or incandescent light may also bring in large numbers.
3. Malaise traps. These work very well to sample many species.
4. CO2 trap. These traps collect females of those species that feed on vertebrate hosts (generally only those of species of Culicoides). A serious problem is that males are not collected and therefore cannot be studied.
Larvae and pupae are obviously restricted to specific habitats and may be sampled directly from those aquatic and semiaquatic habitats where they are found. Reading the section “How and Where do Biting Midges Live?” will introduce the reader to the variety of habitats in which biting midge larvae and pupae may be found. Here are some good sampling methods:
1. Samples of substrate from aquatic habitats (e.g. mud, silt, detritus, leaves) placed in a pan and with water added. Most Ceratopogoninae larvae swim with a rapid serpentine motion and these may be removed with an eyedropper. Pupae almost always float at the water surface and can be taken from the margins of ponds and streams (see Borkent 2014 for more info on pupae). Some workers add salt or sugar to float larvae and pupae from such samples and this is the only way to collect those few species in which the larvae do not swim or the pupae do not float at the surface.
2. Berlese funnels are a good method for extracting larvae from semiaquatic or merely moist material (e.g. mosses growing on trees in cloud forest).
Generally specimens should be preserved in 70% alcohol (or some other preservative). Vials should have air bubbles removed (as much as possible) because sloshing around during transportation from the field to the lab (or home) can break specimens. All genera can be identified while in alcohol with a dissecting microscope.
Studying biting midges requires some equipment beyond what is needed for collecting, slide mounting (see below) and pinning. While in the field, 2-4X reading glasses allow the student to peer more closely at the specimens at hand. While preserved in a glass vial of ethanol in the field a hand lens (15X) is useful for identifying adults and some pupae to genus (once skills have been acquired in the lab using the keys) and larvae to at least subfamily. Once back in the laboratory (or your kitchen table), specimens can be examined in a petri dish of ethanol under a dissecting microscope with good optics of up to at least 40X magnification and strong light. This would allow many of the species here to be identified to at least genus and many to species, depending on the region you live in. I use a dissecting Wild M3 microscope with 15X eyepiece and a 1.5 objective that gives 100X magnification with good resolution and this enhances examination of fine details of the smaller Ceratopogonidae – for sorting larger samples to genus and morphospecies, 40X is often superior, giving a wider field to view. A good, strong light source is also important when examining specimens with a dissecting microscope.
For species in some genera, it is necessary to examine at least the male genitalia mounted on a slide with a compound microscope and for the particularly small specimens (e.g. some Dasyhelea, Brachypogon, some Culicoides) a magnification of 250X is necessary. If investing in new (or second-hand) microscopes it is important to note that some makes claim high magnification have poor optics with poor resolution – a fuzzy view, even if large, is still fuzzy. Some good deals on microscopes are sometimes present on Ebay.
Specimens to be identified to species often need to be placed on microscope slides to be identified (depending on where you live). Methods vary but the following gives excellent results for slide mounting adult specimens stored in ethyl alcohol and which are not too old (up to about 7 years or, if stored under dark and cold conditions, sometimes up to 15 years). This method works particularly well for material to be used for taxonomic studies. A more rapid method is given below.
Prepare for slide mounting by putting the following solutions into a series of stender dishes: 15% acetic acid, 2-propanol, 2-propanol layered over clove oil, clove oil (each dish about 3/4 full). Each specimen needs to go through this sequence of solutions. I prepare 5 specimens at a time, so that I use 20 dishes in total at a time. Put all the dishes on a thin board so that they can be moved around as a group on the table top; this makes it easier to look at specimens while they are in the solutions and still put all the dishes out of harm’s way while they are soaking.
A series of slides needs to be ready, by cleaning these well with a cloth or paper towel and placing a blank slide label on each slide. These slides should be placed in a slide tray. Write the locality number and specimen number on the first five slides to correspond to the first five specimens which are being prepared. In making the following preparations, round coverslips which are 10 mm in diameter work very well.
1. Put specimen into a dish of ethyl alcohol under the microscope. Remove the wings by grabbing the very base of the wing with forceps and pulling it off. Place both wings in the dish with 15% acetic acid.
2.For most specimens, separate head and abdomen from thorax. For very small specimens, just tear the membrane between the head and thorax and between the abdomen and thorax so that all the parts remain attached to each other.
3. Place parts, or torn specimen, into a 3 dram vial which is filled about 1/4 to 1/3 with 8% KOH. Prepare a total of 5 specimens in this way. Each vial needs to be numbered with a grease pencil so that it can be associated with the wings that are in the acetic acid. Place vials of KOH into a beaker partially filled with water. Place beaker on hot plate and heat water to boiling point. Specimens will clear (muscles will disappear) after 2-5 minutes, depending on the size of the specimens. It is important that specimens of a similar size are “cooked” at the same time so that they all clear at the same rate.
4. The following step requires some speed because specimens are damaged by being in hot KOH for too long. This is what needs to be done: remove the specimen from the vial into a dish with some 8% KOH in it and put under the microscope to see if it is properly cleared. The muscles should have dissolved and freely flow out of the specimen if squeezed very, very gently with the forceps (use this method only until you become familiar with what to look for; I never do this anymore). Also, if the specimens are over-cleared they will make for a poor microscope preparation. If cleared sufficiently, place specimen in acetic acid along with the associated wings. Leave for 15 minutes (or longer if necessary).
5. Move specimen to 2-propanol for 15 minutes (or longer if necessary).
6. Move specimen to 2-propanol layered over clove oil. Leave till specimen has sunk to the bottom of the dish which generally takes about 20-60 minutes, depending on the size of the specimen (it can be left longer if necessary). After a day or two, the 2-propanol will mix with the clove oil and some more 2-propanol should be added so that there is always a distinct layer between the 2-propanol and the clove oil. If the clove oil has too much 2-propanol mixed into it, leave the dish without a lid for a few hours and the 2-propanol will evaporate from the mix.
7. Move specimen to pure clove oil and leave for at least 30 minutes (or longer if necessary).
8.Place small drop of Canada Balsam on the microscope slide and place wings in drop. Put coverslip over wings. If Canada Balsam has already hardened too much to move the wings around easily, add a small drop of xylene, which will make the Canada Balsam more fluid. Use xylene for the following procedures as well; it is always needed because Canada Balsam dries faster than the time it takes for a good slide to be made.
9.Place three small drops of Canada Balsam on the microscope slide and put the head, thorax and abdomen into each. If the head, thorax and abdomen are still partially attached (small specimens), put specimen into the drop which will contain just the thorax and remove the head and abdomen; put the head and abdomen into their proper places.
10. Remove the right legs and antennae and place all these under a separate coverslip.
11. Arrange the thorax so that the left side is facing upward. Do the same for the head.
12.Details of the genitalia are often very important in species identification, so that this step is a very important one. Arrange the abdomen so that the ventral part of the genitalia is facing upward. In some males, the genitalia may be only partially turned in relation to the rest of the abdomen and in this case, sometimes the genitalia will need to be removed from the abdomen (by tearing with the forceps and a pin) so that the separated genitalia lies with the ventral side pointed up. The position of the rest of the abdomen is not important. Put sufficient Canada Balsam onto the genitalia and gently put a coverslip over the abdomen. By pushing the coverslip around it is most often possible to ensure that the genitalia are lying in the proper position. With females, it sometimes helps to push the coverslip down very slightly so that the abdomen is slightly flattened and kept in the correct position.
13. Put the slide into the drying oven at 40-50%C. After 1-2 days, the Canada Balsam with the head, abdomen and thorax will have dried somewhat. Add some more Canada Balsam to each of these and very gently put a coverslip over each, so they are not crushed. The partially dried Canada Balsam should keep these parts from being crushed. While the slide is drying, some of the Canada Balsam may leave a small air bubble under the coverslip. If so, add a tiny bit of Canada Balsam. Don’t worry if the added Canada Balsam traps a bubble under the coverslip; the bubble nearly also moves out by itself (after some time).
14. Put the slide back into the drying oven. Check for air bubbles for the next several days and add Canada Balsam if needed (again, don’t worry about trapped bubbles). Remove the slide after 1-2 weeks and put into a slide box. Keep the slide box upright, so the slides are still horizontal.
A group of five specimens can be done at one time (dissection, KOH, acetic acid). After these are all in 2-propanol, a second group of five may be prepared, so that several series of five specimens may be in process at the same time, one group of five following the previous group of five.
Larvae and pupae may be treated with the same solutions and should be mounted dorsal side up. Larvae require no dissection but the operculum of the pupa as well as one of the respiratory organs should be remove and placed under a separate coverslip. If the larva or pupa is represented by an exuviae, the specimens obviously do not require clearing and may be placed directly into 2-propanol and processed from there.
A simpler method for mounting specimens, which is particularly useful for mass mounting material which is likely to include previously described species, is as follows: remove specimens from 70% ethanol (many can be done as one lot), very briefly blot excess alcohol, transfer to 100% ethanol saturated with phenol, leave overnight or longer, and mount specimens on microscope slides in a one to one mixture of Canada Balsam / phenol-ethanol solution. Depending on the stage, specimens may be dissected on the slide and one or more coverslips be placed on the parts. As the phenol-ethanol solution evaporates during drying, more Canada Balsam needs to be added. This method provides good specimens but does not remove internal tissues making the observation of some structures of some taxa difficult to see clearly. Workers using this method should use a fume hood.