The laboratory diagnosis of parasitism


While this technique will detect the eggs and larvae of most nematodes, cestodes and coccidia, it will not demonstrate trematode eggs, which have a higher specific density. For these, a flotation fluid of higher specific gravity such as a saturated solution of zinc sulphate has to be used or a sedimentation method employed as described below.


SEDIMENTATION METHODS


For trematode eggs:



1. Homogenise 3 g of faeces with water and pass the suspension through a coarse mesh sieve (250 μm). Thoroughly wash the material that is retained on this screen, using a fine water jet and discard the debris.

2. Transfer the filtrate to a conical flask and allow to stand for 2 minutes, remove the supernatant, and transfer the remainder (approximately 12–15 ml) to a flat-bottomed tube.

3. After sedimentation for a further 2 minutes the supernatant is again drawn off, a few drops of 5% methylene blue added and the sediment screened using a low power stereomicroscope. Any trematode eggs are readily visible against the pale blue background.

For lungworm larvae, the Baerman apparatus may be used. This consists of a glass funnel held in a retort stand. A rubber tube, attached to the bottom of the funnel, is constricted by a clip. A sieve (aperture 250 μm) is placed in the wide neck of the funnel, which has been partially filled with water, and a double layer of gauze is placed on top of the sieve. Faeces are placed on the gauze and the funnel is slowly filled with water until the faeces are immersed. Alternatively, faeces are spread on a filter paper, which is then inverted and placed on the sieve. The apparatus is left overnight at room temperature during which the larvae migrate out of the faeces and through the sieve to sediment in the neck of the funnel. The clip on the rubber is then removed and the water in the neck of the funnel collected in a small beaker for microscopic examination in a petri dish.


A simple adaptation of the above method is to suspend the faeces enclosed in gauze in a wine glass filled with water and leave overnight. The larvae will leave the faeces, migrate through the gauze and settle at the bottom of the glass. After siphoning off the supernatant, the sediment is examined under the low power of the microscope as above.


CULTURE AND IDENTIFICATION OF LARVAE


Two techniques are widely used for the culture of infective larvae from nematode eggs.


In the first, faeces are placed in a jar with a lid and stored in the dark at a temperature of 21–24°C. The lid should be lined with moist filter paper and should not be tightly attached. After 7 days’ incubation, the jar is filled with water and allowed to stand for 2–3 hours. The larvae will migrate into the water and the latter is poured into a cylinder for sedimentation. The larval suspension can be cleaned and concentrated by using the Baermann apparatus as described above and then killed by adding a few drops of Lugol’s iodine and examined microscopically.


An alternative method is to spread faeces on the middle third of filter paper placed in a moistened petri dish. After storage at 21–24°C for 7–10 days, the dish is flooded with water and the larvae harvested as before.


The identification of infective larvae is a specialist technique and requires some experience. A key to the identification of infective larvae is provided in Table 15.4 and Figure 15.12.


RECOVERY OF ALIMENTARY NEMATODES


Details are given below of a technique for the collection, counting and identification of the alimentary nematodes of ruminants. The procedure is similar for other host species, information on identification being available in the text.



1. As soon as possible after removing the alimentary tract from the body cavity, the abomasal/duodenal junction should be ligatured to prevent transfer of parasites from one site to the other.

2. Separate the abomasum, small intestine and large intestine.

3. Open the abomasum along the side of the greater curvature, wash contents into a bucket under running water and make the total volume up to 2–4 litres.

4. After thorough mixing transfer duplicate 200 ml samples to suitably labelled containers and preserve in 10% formalin.

5. Scrape off the abomasal mucosa and digest in a pepsin/HCl mixture at 42°C for 6 hours; 200 g of mucosa will require 1 litre of mixture. Make digest up to a volume of 2 or 4 litres with cold water and again take duplicate 200 ml samples. Alternatively, the Williams technique may be used. In this, the washed abomasum is placed, mucosal surface down, in a bucket containing several litres of normal saline and maintained at 40°C for 4 hours. Subsequently, the abomasum is gently rubbed in a second bucket of warm saline. The saline from both buckets is poured through a sieve (aperture 38 μm, about 600 to 1 inch) and the residue examined.

6. Open the small intestine along its entire length and wash contents into a bucket. Treat as for the abomasal contents, but digestion of mucosal scrapings is unnecessary.

7. The contents of the large intestine are washed into a bucket, passed through a coarse mesh sieve (aperture 2–3 mm) and any parasites present collected and formalised.

WORM COUNTING PROCEDURE



1. Add 2–3 ml of iodine solution to one of the 200 ml samples.

2. After thorough mixing, transfer 4 ml of suspension to a petri dish, scored with lines to facilitate counting; add 2–3 ml sodium thiosulphate solution to decolourise debris. If necessary, worms may be preserved in an aqueous solution of 10% formalin or 70% alcohol. To clear large worms for microscopic examination, immerse in lactophenol for a suitable period prior to examination.

3. Examine for the presence of parasites using a stereoscopic microscope (×12 objective) and identify and count parasites as male, female and larval stages.

PREPARATION OF SOLUTIONS



  • Pepsin/hydrochloric acid (HCl): dissolve 80 g of pepsin powder in 3 litres of cold water. Add 240 ml concentrated HCl slowly and stir well. Make final volume up to 8 litres. Store at 4°C.
  • Iodine solution: dissolve 907 g of potassium iodide in 650 ml boiling water. Add 510 g iodine crystals and make up to 1 litre.
  • Sodium thiosulphate solution: dissolve 100 g of sodium thiosulphate in 5 litres of water.

KEY TO THE IDENTIFICATION OF GASTROINTESTINAL NEMATODES OF RUMINANTS


Based on the characters described in Tables 15.1(a–c), the following key can be used to differentiate microscopically the genera of some common gastrointestinal nematodes of ruminants.


Body composed of a long filamentous anterior and a short broad posterior region……………………. Trichuris


Body not so divided, oesophagus approximately one third of body length……………………. Strongyloides


Short oesophagus and buccal capsule rudimentary …………………………………… Trichostrongyloidea (A)


Short oesophagus and buccal capsule well developed ………………………………………………… Strongyloidea (B)


(A) TRICHOSTRONGYLOIDEA



1. Distinct cephalic vesicle. Spicules very long uniting in a membrane at the tip………….. Nematodirus Cephalic vesicle small. Spicules relatively short and unjoined posteriorly……………………. Cooperia

2. No cephalic vesicle. Excretory notch present in both sexes…………………………………….. Trichostrongylus Absence of excretory notch……………………. 3

3. Dorsal lobe of bursa asymmetrical, barbed spicules. Large prominent vulval flap in female ………………………………………….. Haemonchus Dorsal lobe of bursa is symmetrical. Vulval flap small or absent……………………………….. Ostertagia

(B) STRONGYLOIDEA



4. Buccal capsule cylindrical……. Oesophagostomum Buccal capsule well developed……………………. 5

5. Slight dorsal curvature of head and presence of cutting plates…………………………….. Bunostomum Absence of teeth, rudimentary leaf crowns present ………………………………………………… Chabertia

Table 15.1 Guide to adult alimentary nematodes of ruminants.


Table 15.1(a) Abomasal worms.


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Table 15.1(b) Small intestinal worms.


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Table 15.1(c) Large intestinal worms.


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IDENTIFICATION OF NEMATODE EGGS


The presence of nematode eggs in faeces is a useful aid to diagnosis of worm infections as they can be identified and counted in faecal samples (Figs 15.215.10). Strongyle eggs are approximately 60–80 μm long, oval, thin-shelled, contain 4–16 cells and are not easily differentiated; but eggs of Trichuris, Nematodirus spp and Strongyloides can be identified and may be counted and reported separately.


THIRD-STAGE LARVAL IDENTIFICATION


It is often useful to know whether faecal egg counts (FECs) are dominated by worms of one particular genus or not, particularly on farms where Haemonchus occurs. If so, larval culture and differentiation can be performed, usually using the faeces from the FEC. This technique takes a further 10–14 days, so results are not available for some time after the FEC is known.


Fig. 15.2 Nematode eggs.


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Table 15.2 Cattle worm egg counts – guide to interpretation.


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Larval differentiation (Table 15.4,Figs 15.11 and 15.12) requires the hatching of the eggs in the sample, culture in sterile faeces or peat, and the subsequent identification of the larvae. Usually, 50 or 100 larvae are counted, and the percentage of each genus reported. It should be noted that eggs of each genus do not always hatch at the same rate because of differences in temperature requirements for the different genera. Larval culture results should therefore be used as a general indication of the worm genera present, rather than a precise determination of the proportion of the FEC contributed by each genus.


Larvae can be identified in a similar manner for pasture samples (see later).


TECHNIQUE


A small drop of suspension of larvae is placed on a microscope and a drop of Gram’s iodine added and a coverslip placed over the drops. The iodine kills the larvae and allows for easier identification of the salient features (Fig. 15.12).


RECOVERY OF LUNGWORMS


For Dictyocaulus, this is best done by opening the air passages starting from the trachea and cutting down to the small bronchi with fine, blunt-pointed scissors. Visible worms are then removed from the opened lungs and transferred to glass beakers containing saline. The worms are best counted immediately, failing which they should be left overnight at 4°C which will reduce clumping. Additional worms may be recovered if the opened lungs are soaked in warm saline overnight.


Another method is Inderbitzen’s modification of the perfusion technique, described by Wolff et al. (1969), in which the lungs are perfused, is as follows. The pericardial sac is incised and reflected to expose the pulmonary artery in which a 2 cm incision is made. Rubber tubing is introduced into the artery and fixed in situ by double ligatures. The remaining large blood vessels are tied off and water from a mains supply allowed to enter the pulmonary artery. The water ruptures the alveolar and bronchiolar walls, flushes out the bronchial lumina, and is expelled from the trachea. The fluid is collected and its contents concentrated by passing through a fine sieve (aperture 38 μm). As before, this is best examined immediately for the presence of adult worms and larvae.


The smaller genera of lungworms of small ruminants are difficult to recover and enumerate, although the Inderbitzen technique may be of value.


Table 15.3 Sheep worm egg counts – guide to interpretation.


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Fig. 15.3 Worm eggs from ruminants.


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Fig. 15.4 Worm eggs from horses.


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Fig. 15.5 Worm eggs from pigs.


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Fig. 15.6 Worm eggs from dogs and cats.


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Fig. 15.7 Worm eggs from poultry.


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Fig. 15.8 Worm eggs from rabbits.


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Fig. 15.9 Worm eggs from rodents.


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Fig. 15.10 Worm eggs from reptiles.


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Table 15.4 Key characteristics used in the identification of third-stage larvae (see Figs 15.11 and 15.12).


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Fig. 15.11 Third-stage larva.


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Fig. 15.12 Key to the identification of third-stage larvae of sheep gastrointestinal nematades. A, Teladorsagia circumcincta; B, Trichostrongylus spp; C, Haemonchus contortus; D, Cooperia spp; E, Nematodirus: (a) battus, (b) filicollis, (c) spathiger; F, Oesophagostomum spp.


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RECOVERY OF TREMATODE AND CESTODE PARASITES


For both Fasciola and Dicrocoelium the livers are removed and cut into slices approximately 1 cm thick. On squeezing the liver slices, any flukes seen grossly are removed and formalised and the slices immersed in warm water overnight. The gallbladder should also be opened and washed, and any flukes removed.


After soaking, the liver slices are again squeezed, rinsed in clean water and discarded. Both washings are passed through a fine sieve (aperture 100 μm) and the material retained and formalised. In the case of intestinal paramphistomes, the first 4 m of duodenum should be tied off, opened, washed and examined for adherent trematodes.


Counts are carried out microscopically, entire flukes plus the numbers of heads and tails being recorded. The highest number of either of the latter is added to the number of entire flukes to give the total count.


Cestodes are usually readily visible in the intestine or liver, but whenever possible these should be removed intact so that, if necessary, the head and the mature and gravid segments are all available for specialist examination. In the case of Echinococcus in canids, however, the worms are so small that the more detailed examination described in the text should be undertaken.


OTHER AIDS TO DIAGNOSIS


There are two other techniques which are useful aids in the diagnosis of trichostrongyle infections in ruminants. The first is the plasma pepsinogen test and the second the estimation of infective larvae on herbage. Both of these techniques should be undertaken in a specialist parasitology laboratory, but a short account is given here of the material required for these tests, the basis of the techniques and how the results may be interpreted.


THE PLASMA PEPSINOGEN TEST


The estimation of circulating pepsinogen is of value in the diagnosis of abomasal damage, and is especially elevated in cases of ostertagiosis. Elevations also occur with other gastric parasites such as Trichostrongylus axei, Haemonchus contortus and, in the pig, Hyostrongylus rubidus.


The principle of the test, which is best carried out by a diagnostic laboratory, is that the sample of serum or plasma is acidified to pH 2.0, thus activating the inactive zymogen, pepsinogen, to the active proteolytic enzyme pepsin. This activated pepsin is then allowed to react with a protein substrate (usually bovine serum albumin) and the enzyme concentration calculated in international units (μmol tyrosine released per 100 ml serum per minute). The tyrosine liberated from the protein substrate by the pepsin is estimated by the blue colour, which is formed when phenolic compounds react with Folin–Ciocalteu’s reagent. The minimum requirement for the test, as carried out in most laboratories, is 1.5 ml serum or plasma. The anticoagulant used for plasma samples is either EDTA or heparin.


In parasitic gastritis of ruminants due to Ostertagia spp and T. axei the levels of plasma pepsinogen become elevated. In parasite-free animals the level is less than 1.0 iu of tyrosine; in moderately infected animals, it is between 1.0 and 2.0 and in heavily infected animals it usually exceeds 3.0, reaching as high as 10.0 or more on occasion. Interpretation is simple in animals during their first 18 months, but thereafter becomes difficult as the level may become elevated when older and immune animals are under challenge. In such cases the absence of the classical clinical signs of diarrhoea and weight loss indicates that there are few adult parasites present.


PASTURE LARVAL COUNTS


For this technique, samples of grass are plucked from the pasture and placed in a polythene bag, which is then sealed and dispatched to a laboratory for processing. It is important to take a reasonable number of random samples, and one method is to traverse the pasture and remove four grass samples at intervals of about four paces until approximately 400 have been collected (Figure 15.13). Another, primarily for lungworm larvae, is to collect a similar number of samples from the close proximity of faecal pats. At the laboratory, the grass is thoroughly soaked, washed and dried and the washings containing the larvae passed through a sieve (aperture 38 μm; 600 to 1 inch) to remove fine debris. The material retained on the sieve is then Baermanised and the infective larvae are identified and counted microscopically under the high power. The numbers present are expressed as L3 per kg of dried herbage.


Fig. 15.13 Butterfly route.


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Where counts in excess of 1000 L3/kg of ruminant gastrointestinal trichostrongyles are recorded, the pasture can be regarded as moderately infective and values of over 5000 L3/kg can be expected to produce clinical disease in young animals during their first season at grass.


Although this is a useful technique for detecting the level of gastrointestinal nematode L3 on pastures, it is less valuable for detecting lungworm larvae because of the rapid fluctuations of these larvae on pastures.


A more sophisticated technique, the Jorgensen method, which depends on migration of larvae through an agar medium containing bile, is used in some laboratories for estimating Dictyocaulus larval populations on pasture; since most lungworm larvae are concentrated close to faeces, herbage samples should be collected from around faecal deposits. In the present state of knowledge, the detection of any lungworm larvae in herbage samples should be regarded with suspicion and even a negative finding does not necessarily imply that the pasture is free of infection.


ECTOPARASITES


Arthropods of veterinary interest are divided into two major groups, the Insecta and the Arachnida. Most are temporary or permanent ectoparasites, found either in or on the skin, with the exception of some flies whose larval stages may be found in the somatic tissues of the host. Parasitic insects include flies, lice and fleas, while the two groups of arachnids of veterinary importance are the ticks and mites. In all cases diagnosis of infection depends on the collection and identification of the parasite(s) concerned.


INSECTS


FLIES


Adult dipteran flies visiting animals are usually caught either by netting or after being killed by insecticides, while larvae may be collected in areas where animals are housed or directly from animals where the larval stages are parasitic. Identification of the common flies of veterinary interest, at least to generic level, is fairly simple, the key characters being described in the guide below. Identification of larvae to generic and species level is rather more specialised and depends on examination of certain features such as the structure of the posterior spiracles. Publications dealing with this may be found in References and further reading.


Guide to the families of adult Diptera of veterinary importance


1 Insects with one pair of wings on the mesothorax and a pair of club-like halteres on the metathorax …………………………………………………………………..2 Wingless insects; may be with or without halteres; body clearly divided into head, thorax and abdomen; three pairs of legs; dorsoventrally flattened; brown in colour; 5–8 mm in length; resident on sheep, horses, deer, goats or wild birds (Fig. 15.14)………………………………………….5

2 Antennae composed of three segments; third segment usually with an arista; foot with two pads (Fig. 15.15) …………………………………………. ……………………………………………..Cyclorrhapha 3 Antennae composed of three sections; third antennal section enlarged and composed of four to eight segments; palps two-jointed with the second segment enlarged; foot with three pads; vein R4+5 forks to form a large ‘Y’ across the wing tip (Fig. 15.16); large, stout bodied flies with large eyes……………………………..Tabanidae 12 Antennae long, slender and composed of many articulating segments; palps composed of four to five segments; small slender flies with long narrow wings…………………………….Nematocera 13

Fig. 15.14 Adult sheep ked, Melophagus ovinus.


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3 Frons with ptilinal suture (Fig. 15.17)………………. …………………………………….Series Schizophora 4 Frons without ptilinal suture………..Series Aschiza

4 Second antennal segment usually with a groove (Fig. 15.17); thoracic transverse suture strong (Fig. 15.19); thoracic squamae usually well developed (Fig. 15.20)…………………………..Calypterae 5 Second antennal segment usually without a groove; thoracic transverse suture weak; thoracic squamae often vestigial…………………….Acalypterae

5 Thorax broad and dorsoventrally flattened; may appear spider or tick-like; often wingless (Fig. 15.14); wings when present with venation abnormal with veins crowded into leading half of wing…………………………………………Hippoboscidae Wings with veins not crowded together towards the leading edge; thorax not dorsoventrally flattened……………………………………………………….6

Fig. 15.15 Variations in the antennae found in the three suborders of Diptera.


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Fig. 15.16 Antennae of (a) Chrysops, (b) Haematopota and (c) Tabanus. (d) Wing venation of Tabanidae (reproduced from Smart, 1943).


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Fig. 15.17 The principal features of the dichoptic head of a typical adult Calypterate cyclorrhaphous dipteran (redrawn from Smart, 1943).


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Fig. 15.18 Wing venation typical of species of Glossina, showing the characteristic hatchet shape of the cell dm.


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Fig. 15.19 The principal features of the generalised thorax of an adult Calypterate cyclorrhaphous dipteran. (a) Dorsal view. (b) Lateral view (redrawn from Smart, 1943).


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Fig. 15.20 The veins and cells of the wings of a typical calypterate dipteran, Calliphora vicina.


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6 Proboscis long, forwardly directed and embraced by long palps; arista with feathery short hairs present only on dorsal side; discal medial cell of wings characteristically ‘hatchet’ shaped (Fig. 15.18); found only in sub-Saharan Africa …………………………………………………. Glossinidae Discal medial cell of wings widening gradually and more or less regularly from the base………….7

7 Mouthparts small, usually functionless; head bulbous; antennae small; flies more or less covered with soft hair; larval parasites of vertebrates……. ………………………………………………………Oestridae Mouthparts usually well developed; antennae not small; flies with strong bristles…………………8

8 Hypopleural bristles present (Fig. 15.19)………….9 Hypopleural bristles absent…………………………..11

9 Post-scutellum strongly developed; larval parasitoids of insects…………………………….Tachinidae Post-scutellum weak or absent………………………10

10 Dull grey appearance; three black stripes on the scutum; abdomen usually with chequered or spotted pattern (Fig. 15.21); larval parasites of vertebrates………………………………..Sarcophagidae Metallic, iridescent appearance (blue–black, violet– blue, green); larval parasites of vertebrates…………. …………………………………………………..Calliphoridae

11 Wings with vein A1 not reaching the wing edge; strong curved A2 vein the tip of which approaches A1 (Fig. 15.22); aristae bare…………… …………………………………………………….. Fanniidae Wings with vein A1 not reaching the wing edge; A2 vein not strongly curved (Fig. 15.23); aristae bilaterally plumose to the tip……………….Muscidae

12 Antennal flagellum with four segments (Fig. 15.16); wings mottled; proboscis shorter than head …………………………..Haematopota (Tabanidae) Antennal flagellum with five segments (Fig. 15.16); apical spurs on tibiae are small and may be hidden by hair; wings usually with costal region dark and a single dark broad transverse band; proboscis shorter than head……………………. ……………………………………..Chrysops (Tabanidae) Antennal flagellum with five segments (Fig. 15.16); no apical spurs on hind tibiae; wings usually clear but may be dark or banded; proboscis shorter than head………………Tabanus (Tabanidae)

13 Small, hairy, moth-like flies; numerous parallel wing veins running to the margin; wings pointed at the tip…………………………………..Psychodidae 14 Not like this………………………………………….15

14 Palps five-segmented; biting mouthparts at least as long as head; antennal segments almost cylindrical; two longitudinal wing veins between radial and medial forks (Fig. 15.24)………………………….. ………………………………………………. Phlebotominae

15 Ten or more veins reaching the wing margin ……………………………………………………………….. 16 Not more than eight veins reaching the wing margin……………………………………………………….17

16 Wing veins and hind margins of wings covered by scales (Fig. 15.25); conspicuous forward-projecting proboscis…………………………………………. Culicidae

17 Wings broad; wing veins thickened at the anterior margin; antennae not hairy; thorax humped; antennae usually with 11 rounded segments; palps long with five segments extending beyond the proboscis; first abdominal tergite with a prominent basal scale fringed with hairs (Fig. 15.26) …………. …………………………………………………….. Simuliidae Wings not particularly broad, antennae hairy ……. ……………………………………………………………….. 18

18 Front legs often longer than others; median vein not forked; (non-biting midges) …………………….. ………………………………………….Chironomidae Front legs not longer than others; wings with median vein forked; antennae with 14–15 visible segments; palps with five segments; female mouthparts short; legs short and stout; two radial cells and cross vein r-m strongly angled in relation to media; at rest wings close flat over abdomen (Fig. 15.27)…………………………………. ………………………………Culicoides (Ceratopogonidae)

Fig. 15.21 (a) Adult of the flesh fly Sarcophaga carnaria (reproduced from Castellani and Chalmers, 1910). (b) Wohlfahrtia magnifica, abdomen of adult (reproduced from Smart, 1943).


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Fig. 15.22 Wing venation typical of species of Fannia, showing the characteristic convergence of the anal veins.


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Fig. 15.23 (a) Female house fly, Musca domestica. (b) Wing venation typical of species of Musca, showing the strongly bent vein M ending close to R4+5 (after Smart, 1943).


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Fig. 15.24 (a) Adult female sandfly, Phlebotomus papatasi. (b) Wing venation typical of species of Phlebotomus (Psychodidae) (reproduced from Smart, 1943)


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Fig. 15.25 Aedes atropalpus: adult (reproduced from Eidmann and Kuhlhorn, 1970).


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Fig. 15.26 (a) Adult female Simulium. (b) Wing venation typical of Simulium, showing the large anal lobe and crowding of the veins towards the leading edge (reproduced from Smart, 1943).


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Fig. 15.27 (a) Adult female Culicoides nebeculosus at rest. (b) Wing venation typical of species of Culicoides, showing the two elongate radial cells (reproduced from Edwards et al., 1939).


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Guide to third-stage larvae causing myiasis in domestic animals

The guide to larvae presented below applies specifically to recognition of the third stage. This stage is usually of the longest duration and, since the larvae are approaching their maximum size or are beginning to wander, is usually the stage when they are most commonly observed. It should be noted, that because the external structure of larvae may change over the course of their growth and development, first- and second-stage larvae may not key out appropriately.



1 Body more or less cylindrical; no obvious head capsule…………………………………………………….. 2 Fly larvae with an obvious head capsule; rarely found associated with livestock myiasis…………… ……………………..Diptera, Nematocera or Brachycera

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